CHEMICAL BIOLOGY

Chemistry of Lipid Domains

 

Richard M. Epand,, McMaster University Health Sciences Centre,, Hamilton, Ontario,, Canada

doi: 10.1002/9780470048672.wecb283

 

Biological membranes are composed largely of proteins and lipids. A wide range of molecular structures exists among these molecules. It is therefore not surprising that biological membranes are not uniform but rather cluster certain molecules in specific regions or domains. This behavior can be mimicked in model systems comprising a limited number of components to study in more detail the molecular nature of this domain formation. Two types of domains exist that have been more extensively studied. One is a membrane domain enriched in polyanionic lipids, such as phosphatidylinositol diphosphate. Such domains are formed by the presence of proteins with segments containing several cationic amino acid residues. Such proteins have been termed ''pipmodulins.'' Another kind of domain is formed as a consequence of the nonuniform distribution of cholesterol in the membrane. Caveolae represent one type of cholesterol-rich domain that is well characterized. Other cholesterol-rich domains are termed ''rafts.'' Characterization of rafts in model membranes is well documented, but the nature of cholesterol-rich domains in biological membranes remains a subject of controversy. Several imaging and fluorescence methods are being employed to further characterize the size and lifetime of small raft domains in biological membranes.

 

There is much current interest in understanding the properties of domains in biological membranes. It is well established that these membranes are not homogeneous with uniform composition and properties throughout. However, a continual gradation exists between transient, nonuniform clustering of a small number of molecules and the formation of a large or more stable cluster. The former might be better described as the nonideal mixing of components, whereas the latter is more appropriate to be called a domain. No quantitative criteria exist to differentiate between these two extents of nonideal mixing. In this article, we will discuss the possible biological roles of membrane domains, the nature of the domains that exist in biological membranes, and the driving force for their formation. In addition, we will discuss some methods for studying membrane domains and how these methods have contributed to our current state of knowledge of this phenomenon.

 

Biological Background

To begin with, biological membranes are not uniform with respect to membrane sidedness. Both the protein and the lipid composition of these membranes exhibit transbilayer asymmetry. In addition, in polarized mammalian cells, a segregation of both protein and lipid molecules exists between the apical and basolateral surfaces (1-4). Both transbilayer asymmetry and segregation in polarized cells occur over a long time period with slow exchange between domains. In addition, smaller, more transient domains are present along the plane of the membrane. The roles of these transient lateral domains are likely to be the clustering of molecules in the small volume of a membrane domain to allow for their efficient interaction, which may be particularly important for signal transduction processes in which a series of chemical reactions can lead to amplification of a signal. In addition, the nonuniform distribution of lipid components will result in the physical properties of the membrane varying between a domain and the bulk of the membrane, which, in turn, will modulate the activity of membrane proteins.

 

Chemistry

We will discuss the chemical composition, size, and lifetime of domains in biological and model membranes. This area is currently under very active investigation, and we are likely to see further developments in the future. We will focus particularly on two types of membrane domains for which there is more evidence for their existence in biological membranes and more characterization of their properties. We can classify these domains as either cholesterol-rich domains or anionic lipid clusters.

 

Cholesterol-Rich Domains

Cholesterol is a major lipid component of mammalian cell plasma membranes, accounting for approximately 35% of the total lipid of the membrane. Cholesterol has a chemical structure that is very different from the major polar lipid constituents, which is a consequence of the fused ring system of cholesterol that gives this lipid less conformational flexibility than the straight chains of the polar lipids. It is thus not surprising that cholesterol does not mix well in membranes and that cholesterol segregates as crystals at around 50-60 mol% in bilayers of several lipids and at a much lower mol fraction of cholesterol in bilayers comprising lipids with unsaturated acyl chains (5).

The lateral distribution of molecules in a membrane will be the result of differences in the interaction energies between different groups of molecules. As the chemical structure of cholesterol is very different from that of polar lipids, differences will likely exist in the stability of complexes of peptides or proteins with cholesterol versus those with phospholipids. This property will result in cholesterol being distributed nonuniformly in the membrane at thermodynamic equilibrium. In addition, cholesterol will have different extents of interaction with saturated versus unsaturated polar lipids, a well-studied example of which in model liposomes is the fluid-fluid immiscibility resulting in the formation of domains in mixtures of dioleoylphosphatidyl-choline (DOPC), sphingomyelin, and cholesterol. In these mixtures, a saturated acyl chain of sphingomyelin has more favorable interactions with cholesterol than do the unsaturated oleoyl chains. If a phosphatidylcholine with saturated acyl chains, di- palmitoylphosphatidylcholine (DPPC), replaces palmitoyl sphingomyelin (PSM) in mixtures with DOPC and cholesterol, the two phase diagrams are essentially identical (6). The size of the domains formed in such a lipid mixture is of the order of microns in diameter (7, 8). One domain is highly enriched in sphingomyelin, whereas the other contains phosphatidylcholine as the major phospholipid. As indicated above, the cholesterol partitions preferentially into the more ordered sphingomyelin domain, although the segregation of cholesterol between the domains is only partial (9). Interestingly, substituting DOPC with 1-palmitoyl-2-oleoyl phosphatidylcholine (POPC) in these ternary lipid mixtures with cholesterol results in a large difference between the phase diagrams of POPC/cholesterol/PSM and POPC/cholesterol/DPPC with no miscibility transition observed with the latter mixture (6). In addition to differing in chemical composition, the two immiscible domains have different physical properties. The more ordered domain is the one enriched in sphingomyelin. This phase is still a fluid phase, but it has increased order and has been termed a liquid-ordered phase. Specific properties of the liquid-ordered phase place it between a liquid-disordered phase and an ordered phase. The rate of lateral diffusion of the lipid in the liquid-ordered phase is similar to that in a liquid-disordered phase, making it a liquid phase. In contrast, the acyl chain order parameters are closer to that of an ordered or solid phase, which is a consequence of the neighboring cholesterol inhibiting trans to gauche isomerization of the carbon-carbon single bonds. The coexistence of a liquid-ordered and liquid-disordered domain in a mixture of phospholipids and cholesterol will depend on the chemical nature of the phospholipids, the relative proportion of each of the lipid components, and the temperature. The phase diagram of several three-component systems (two phospholipids and cholesterol) in excess water as a function of temperature has been determined.

Although the characterization of membrane domains that depend on the presence of cholesterol have been well studied and largely understood in model systems, the relevance of this behavior to domains in biological membranes is controversial. However, one type of cholesterol-rich domain exists in the plasma membrane of certain mammalian cells for which there is considerable evidence (10): the caveolae. Caveolae have a lipid composition with an enrichment of cholesterol and sphingomyelin (11) and would therefore be expected to form a liquid-ordered domain that is segregated from the surrounding membrane in a liquid-disordered phase. Caveolae have a distinctive morphological structure of a flask-shaped invagination of approximately 65 nm in diameter that can readily be identified with electron microscopy. This characteristic shape is a consequence of this domain containing a high content of the membrane protein caveolin. Caveolae can be isolated after breaking up the cell membrane, without the use of any detergents (12). Caveolae domains are large, stable, and enriched in cholesterol and sphingomyelin. Their lipid composition and physical state are similar to what is expected for membrane rafts, but the size and lifetime of rafts in biological membranes are much less (13).

The term “raft” has been extensively used in the literature to specify a particular kind of domain in a biological membrane, but its use has often evaded a precise definition. We suggest that a raft be defined as a non-caveolae, cholesterol-rich domain in a biological membrane. As this domain is one with a phase boundary separating it from its surroundings, it would be in a liquid-ordered state. A raft domain would be similar to caveolae except for the distinguishing morphology and presence of the protein caveolin in the latter case. Such a raft domain could be visualized by several imaging methods described below, or its presence could be ascertained with spectroscopic methods. This definition of the physical presence of a cholesterol-rich cluster of molecules in a biological membrane is different from rafts that are defined on the basis of detergent insolubility. The membrane fraction that is insoluble in cold detergent and is in a low density fraction upon ultracentrifugation has been suggested to be similar to raft domains in biological membranes; but it is an extrapolation that is increasingly questioned. This fraction should be referred to as the low density, detergent-resistant fraction and not necessarily representing a raft. Detergent resistance is a phenomenological definition, whereas the term raft should be restricted to a physical structure, a domain, in a membrane.

Several additional features of domains exist in biological membranes that are not present in model systems. Two fundamental features of biological membranes that are not present in model membranes are the presence of proteins and the transbilayer asymmetry. In particular, proteins are a major component of plasma membranes, comprising approximately 50% of the weight of the material. They are also important for domain formation, which is most clear in the case of caveolae, whose presence in cells is closely tied to the expression of caveolin (14). These structures can even be induced to form in cells normally lacking caveolae by inducing the expression of caveolin (15), and caveolae are lost in mice lacking caveolin 1 and 3 (16, 17). However, this example is only one that is specific and well-documented of a protein that affects domain formation. In general, any protein that interacts more favorably with some lipids than with others will result in domain formation. It would be anticipated that, as a consequence of cholesterol increasing the packing density of the membrane, many proteins would be excluded from cholesterol-rich regions (18). The preferential interaction of proteins with cholesterol-depleted domains will result in cholesterol concentrating in a different region of the membrane, resulting in the formation of cholesterol-rich domains. An example of such behavior has been shown with an amphipathic helical peptide (19).

 

Electrostatically Driven Domain Formation

Another mechanism for domain formation is through the clustering of anionic lipids by cations or by polycationic protein segments. There has been particular interest in the role of proteins with cationic clusters in causing the formation of domains enriched in the polyanionic phospholipid phosphatidylinosi- tol 4,5-bisphosphate (PIP2) (20). In particular, three proteins have been identified that contain cationic clusters, sequester PIP2, and are largely devoid of structure. These proteins are the myristoylated alanine-rich C-kinase substrate (MARCKS), growth-associated protein 43 (GAP43), and neuronal axonal membrane protein (NAP-22 or CAP23 for the form found in chicken) (21). These proteins have been termed “pipmodulins” because of their ability to modulate the availability of free PIP2. In the case of the MARCKS protein, the electrostatically driven sequestering of PIP2 by the cationic cluster of residues in this protein can be reversed by phosphorylation a Ser residues in this cluster by protein kinase C (22). Another modulation of the sequestering of PIP2 by MARCKS results from the competition between Calcium-calmodulin and the anionic lipid surface for the cluster of basic residues in MARCKS (23, 24).

There has been a suggestion that electrostatically driven formation of PIP2 domains have a relationship to membrane rafts. It is known that several proteins that sequester PIP2 locate in raft domains (25 , 26). In addition, the domain localization of PIP2 is dependent on the presence of cholesterol (27-29). However, the conclusion that PIP2 is localized in raft domains of biological membranes has been questioned (30). In the model system of planar bilayers on a solid support of mica, we showed that a Bodipy-labeled PIP2 partitioned into the liquid-disordered domains (31), which was accomplished using a combination of atomic force microscopy (AFM) and confocal fluorescence microscopy. However, it is known that fluorescence probes can affect the partitioning of fluorescently labeled lipid molecules between membrane domains (32). A more reliable localization of unlabeled PIP2 using antibodies to the lipid component confirmed that the PIP2 located in the lower, liquid-disordered domains as well as in the interfacial region between liquid-ordered and liquid-disordered domains (31).

A few proteins exist that sequester PIP2 in a cholesterol- dependent manner. One of these proteins is the N-terminal myristoylated peptide of NAP-22 (33, 34). Combined confocal microscopy and AFM show that this peptide forms new cholesterol-rich domains within the liquid-disordered domain to which it attracts PIP2 (31). In addition, a peptide segment of caveolin promotes the formation of membrane domains containing both cholesterol and PIP2 (35).

 

Chemical Tools and Techniques

There have been efforts to physically separate membrane domains as well as to determine their properties using imaging, spectroscopic, and calorimetric methods. Several of these methods have been recently reviewed (18). In this article, we will focus on membrane fractionation and on imaging methods.

 

Fractionation

Cholesterol-rich domains of biological membranes have been isolated on the basis of their insolubility in 1% Triton at 4° C (36). This method has come into question because of the possibility that the Triton itself causes rearrangement of membrane components (37, 38). As indicated above, this fraction should not be referred to as a raft. However, the cholesterol-rich domain in the form of caveolae can be isolated without use of detergent (39, 40), providing stronger evidence that this cholesterol-rich domain exists in a biological membrane before extraction.

 

Fluorescence Methods

Fluorescence microscopy has been used extensively to study the formation of membrane domains. It has been particularly successful in determining the phase behavior of model membranes in the form of giant unilamellar vesicles (41). One of the limitations of imaging with light microscopy is that only relatively larger domains can be visualized because the method is limited by the wavelength of light. However, higher resolution can be attained with a near-field scanning optical microscope (NSOM) (42). In addition, recently, the method of stimulated emission depletion has been applied to biological systems that will allow imaging of smaller domains (43). Fluorescent-based methods have also been used to measure the interaction between molecules in small domains of nanometer, rather than micron, size. Fluorescence quenching and fluorescence resonance energy transfer methods have been used to demonstrate the presence of small domains in both model as well as biological membranes (44-46). However, application of these methods to biological membranes has not led to definitive results (45). Recent modeling studies have indicated that the lateral mobility of proteins in membranes is sensitive to the protein dimensions, including protein-lipid interactions (47). Measurements of the lateral diffusion of proteins in the apical plasma membrane of epithelial cells using fluorescence recovery after photobleaching gives evidence for the presence of at least two coexisting liquid phases (2). Another method for the measurement of lateral diffusion in membranes that holds promise for the future is fluorescence correlation spectroscopy (48).

 

Atomic Force Microscopy (AFM)

AFM can readily be applied to bilayers deposited on a solid support. The method has the advantage that it can give nanometer resolution and that it is sensitive to measurements of height. It has been used for studying phase separation in lipid membranes (49). One also has to consider possible interactions between the solid support and the bilayer, particularly with regard to measuring rates of lateral diffusion. One can also use AFM in combination with fluorescence microscopy or with total internal reflection fluorescence (TIRF) microscopy to locate labeled molecules in the membrane. However, one must be cautious about the fluorescent probe affecting the domain localization (32). It is also possible to use force measurements in AFM to determine the extent of the electrical double layer (50) and thereby examine the clustering of charged lipids in a bilayer.

 

Future Directions

Important questions still need to be resolved regarding the nature of the interactions among membrane components that can lead to fluid-fluid immiscibility. However, to extend these model system studies to biological membranes, a deeper understanding of the role of proteins in determining domain formation is required. This task is complicated because there are a large variety of proteins in biological membranes and many motifs by which they can interact with surrounding molecules. Domains can be formed if the proteins interact preferentially with certain membrane components, provided these interactions are sufficiently strong to overcome the entropy of mixing. In addition, a more accurate and complete characterization of domains in biological membranes is required and is an active area of research. These domains are heterogeneous and, therefore, no single description of their size, shape, lifetime, or transbilayer properties exists. Each type of membrane domain will have to be studied in its own right. This complex task will require the coordinated effort of membrane biophysicists, cell biologists, and others. It will likely lead to a better understanding of the coordinated behavior of membrane proteins and the functioning of certain domain-related biological processes such as signal transduction and certain cases of endocytosis and viral fusion.

 

References

1. Fullekrug J, Simons K. Lipid rafts and apical membrane traffic. Ann. NY Acad. Sci. 2004; 1014:164-169.

2. Meder D, Moreno MJ, Verkade P, Vaz WLC, Simons K. Phase coexistence and connectivity in the apical membrane of polarized epithelial cells. PNAS 2006; 103:329-334.

3. Schuck S, Simons K. Polarized sorting in epithelial cells: raft clustering and the biogenesis of the apical membrane. J. Cell Sci. 2004; 117:5955-5964.

4. Simons K, Vaz WLC. Model systems, lipid rafts, and cell membranes. Annu. Rev. Biophys. Biomol. Struct. 2004; 33:269-295.

5. Brzustowicz MR, Cherezov V, Caffrey M, Stillwell W, Wassall SR. Molecular organization of cholesterol in polyunsaturated membranes: microdomain formation. Biophys. J. 2002; 82:285- 298.

6. Veatch SL, Keller SL. Miscibility phase diagrams of giant vesicles containing sphingomyelin. Phys. Rev. Lett. 2005; 94:1481011-1481014.

7. Dietrich C, Bagatolli LA, Volovyk ZN, Thompson NL, Levi M, Jacobson K, Gratton E. Lipid rafts reconstituted in model membranes. Biophys. J. 2001; 80:1417-1428.

8. Veatch SL, Keller SL. Separation of liquid phases in giant vesicles of ternary mixtures of phospholipids and cholesterol. Biophys. J. 2003; 85:3074-3083.

9. Veatch SL, Polozov IV, Gawrisch K, Keller SL. Liquid domains in vesicles investigated by NMR and fluorescence microscopy. Biophys. J. 2004; 86:2910-2922.

10. Parton RG, Hanzal-Bayer M, Hancock JF. Biogenesis of caveolae: a structural model for caveolin-induced domain formation. J. Cell Sci. 2006; 119:787-796.

11. Ortegren U, Karlsson M, Blazic N, Blomqvist M, Nystrom FH, Gustavsson J, Fredman P, Stralfors P. Lipids and glycosphingolipids in caveolae and surrounding plasma membrane of primary rat adipocytes. Eur. J. Biochem. 2004; 271:2028-2036.

12. Westermann M, Leutbecher H, Meyer HW. Membrane structure of caveolae and isolated caveolin-rich vesicles. Histochem. Cell. Biol. 1999; 111:71-81.

13. Hancock JF. Lipid rafts: contentious only from simplistic standpoints. Nat. Rev. Mol. Cell Biol. 2006; 7:456-462.

14. Parton RG. Caveolae and caveolins. Curr. Opin. Cell Biol. 1996; 8:542-548.

15. Fra AM, Williamson E, Simons K, Parton RG. De novo formation of caveolae in lymphocytes by expression of VIP21-caveolin. Proc. Natl. Acad. Sci. U.S.A. 1995; 92:8655-8659.

16. Drab M, Verkade P, Elger M, Kasper M, Lohn M, Lauterbach B, Menne J, Lindschau C, Mende F, Luft FC, Schedl A, Haller H, Kurzchalia TV. Loss of caveolae, vascular dysfunction, and pulmonary defects in caveolin-1 gene-disrupted mice. Science 2001; 293:2449-2452.

17. Galbiati F, Engelman JA, Volonte D, Zhang XL, Minetti C, Li M, Hou H Jr, Kneitz B, Edelmann W, Lisanti MP. Caveolin-3 null mice show a loss of caveolae, changes in the microdomain distribution of the dystrophin-glycoprotein complex, and t-tubule abnormalities. J. Biol. Chem. 2001; 276:21425-21433.

18. Epand RM. Do proteins facilitate the formation of cholesterol-rich domains? Biochim. Biophys. Acta 2004; 1666:227-238.

19. Epand RM, Epand RF, Sayer BG, Melacini G, Palgulachari MN, Segrest JP, Anantharamaiah GM. An apolipoprotein AI mimetic peptide: membrane interactions and the role of cholesterol. Biochemistry 2004; 43:5073-5083.

20. McLaughlin S, Murray D. Plasma membrane phosphoinositide organization by protein electrostatics. Nature 2005; 438:605-611.

21. Laux T, Fukami K, Thelen M, Golub T, Frey D, Caroni P. GAP43, MARCKS, and CAP23 modulate PI(4,5)P(2) at plasmalemmal rafts, and regulate cell cortex actin dynamics through a common mechanism. J. Cell Biol. 2000; 149:1455-1472.

22. Ohmori S, Sakai N, Shirai Y, Yamamoto H, Miyamoto E, Shimizu N, Saito N. Importance of protein kinase C targeting for the phosphorylation of its substrate, myristoylated alanine-rich C-kinase substrate. J. Biol. Chem. 2000; 275:26449-26457.

23. Arbuzova A, Murray D, McLaughlin S. MARCKS, membranes, and calmodulin: kinetics of their interaction. Biochim. Biophys. Acta 1998; 1376:369-379.

24. Kim J, Shishido T, Jiang X, Aderem A, McLaughlin S. Phosphorylation, high ionic strength, and calmodulin reverse the binding of MARCKS to phospholipid vesicles. J. Biol. Chem. 1994; 269:28214-28219.

25. Caroni P. New EMBO members’ review: actin cytoskeleton regulation through modulation of PI(4,5)P(2) rafts. EMBO J. 2001;20:4332-4336.

26. Flesch FM, Yu JW, Lemmon MA, Burger KN. Membrane activity of the phospholipase C-deltal pleckstrin homology (PH) domain. Biochem. J. 2005; 389:435-441.

27. Bodin S, Giuriato S, Ragab J, Humbel BM, Viala C, Vieu C, Chap H, Payrastre B. Production of phosphatidylinositol 3,4,5-trisphosphate and phosphatidic acid in platelet rafts: evidence for a critical role of cholesterol-enriched domains in human platelet activation. Biochemistry 2001; 40:15290-15299.

28. Pike LJ, Miller JM. Cholesterol depletion delocalizes phosphatidylinositol bisphosphate and inhibits hormone-stimulated phosphatidylinositol turnover. J. Biol. Chem. 1998; 273:22298-22304.

29. Rozelle AL, Machesky LM, Yamamoto M, Driessens MH, Insall RH, Roth MG, Luby-Phelps K, Marriott G, Hall A, Yin HL. Phosphatidylinositol 4,5-bisphosphate induces actin-based movement of raft-enriched vesicles through WASP-Arp2/3. Curr. Biol. 2000; 10:311-320.

30. Van Rheenen J, Achame EM, Janssen H, Calafat J, Jalink K. PIP2 signaling in lipid domains: a critical re-evaluation. EMBO J. 2005; 24:1664-1673.

31. Shaw JE, Epand RF, Sinnathamby K, Li Z, Bittman R, Epand RM, Yip CM. Tracking peptide-membrane interactions: insights from in situ coupled confocal-atomic force microscopy imaging of NAP-22 peptide insertion and assembly. J. Struct. Biol. 2006; 155:458-469.

32. Shaw JE, Epand RF, Epand RM, Li Z, Bittman R, Yip CM. Correlated fluorescence-atomic force microscopy of membrane domains: structure of fluorescence probes determines lipid localization. Biophys. J. 2006; 90:2170-2178.

33. Epand RM, Vuong P, Yip CM, Maekawa S, Epand RF. Cholesterol-dependent partitioning of Ptdlns(4,5)P-2 into membrane domains by the N-terminal fragment of NAP-22 (neuronal axonal myristoylated membrane protein of 22 kDa). Biochem. J. 2004; 379:527-532.

34. Epand RF, Sayer BG, Epand RM. Induction of raft-like domains by a myristoylated NAP-22 peptide and its Tyr mutant. FEBS J. 2005; 272:1792-1803.

35. Wanaski SP, Ng BK, Glaser M. Caveolin scaffolding region and the membrane binding region of SRC form lateral membrane domains. Biochemistry 2003; 42:42-56.

36. Brown DA, London E. Functions of lipid rafts in biological membranes. Annu. Rev. Cell Dev. Biol. 1998; 14:111-136.

37. Heerklotz H. Triton promotes domain formation in lipid raft mixtures. Biophys. J. 2002; 83:2693-2701.

38. Lichtenberg D, Goni FM, Heerklotz H. Detergent-resistant membranes should not be identified with membrane rafts. Trends Biochem. Sci. 2005; 30:430-436.

39. Song KS, Li S, Okamoto T, Quilliam LA, Sargiacomo M, Lisanti MP. Co-purification and direct interaction of Ras with caveolin, an integral membrane protein of caveolae microdomains. Detergent-free purification of caveolae microdomains. J. Biol. Chem. 1996; 271:9690-9697.

40. Westermann M, Leutbecher H, Meyer HW. Membrane structure of caveolae and isolated caveolin-rich vesicles. Histochem. Cell Biol. 1999; 111:71-81.

41. Veatch SL, Keller SL. Seeing spots: complex phase behavior in simple membranes. Biochim. Biophys. Acta Mol. Cell Res. 2005; 1746:172-185.

42. Tokumasu F, Hwang J, Dvorak JA. Heterogeneous molecular distribution in supported multicomponent lipid bilayers. Langmuir 2004; 20:614-618.

43. Willig KI, Rizzoli SO, Westphal V, Jahn R, Hell SW. STED microscopy reveals that synaptotagmin remains clustered after synaptic vesicle exocytosis. Nature 2006; 440:935-939.

44. de Almeida RF, Loura LM, Fedorov A, Prieto M. Lipid rafts have different sizes depending on membrane composition: a time-resolved fluorescence resonance energy transfer study. J. Mol. Biol. 2005; 346:1109-1120.

45. Silvius JR, Nabi IR. Fluorescence-quenching and resonance energy transfer studies of lipid microdomains in model and biological membranes. Mol. Membr. Biol. 2006; 23:5-16.

46. Rao M, Mayor S. Use of Forster’s resonance energy transfer microscopy to study lipid rafts. Biochim. Biophys. Acta 2005; 1746:221-233.

47. Gambin Y, Lopez-Esparza R, Reffay M, Sierecki E, Gov NS, Genest M, Hodges RS, Urbach W. Lateral mobility of proteins in liquid membranes revisited. Proc. Natl. Acad. Sci. U.S.A. 2006; 103:2098-2102.

48. Kahya N, Schwille P. Fluorescence correlation studies of lipid domains in model membranes. Mol. Membr. Biol. 2006; 23:29-39.

49. Connell SD, Smith DA. The atomic force microscope as a tool for studying phase separation in lipid membranes. Mol. Membr. Biol. 2006; 23:17-28.

50. Sachs F. Probing the double layer: effect of image forces on AFM. Biophys. J. 2006; 91:L14-L15.

 

Further Reading

Bhat RA, Panstruga R. Lipid rafts in plants. Planta 2005; 223:5-19.

Cheng ZJ, Deep SR, Marks DL, Pagano RE. Membrane microdomains, caveolae, and caveolar endocytosis of sphingolipids. Mol. Membr. Biol. 2006; 23:101-110.

Cohen AW, Hnasko R, Schubert W, Lisanti MP. Role of caveolae and caveolins in health and disease. Physiol. Rev. 2004; 84:1341-1379.

Kenworthy AK. Fleeting glimpses of lipid rafts: how biophysics is being used to track them. J. Investig. Med. 2005; 53:312-317.

Mukherjee S, Maxfield FR. Membrane domains. Annu. Rev. Cell Dev. Biol. 2004; 20:839-866.

Rajendran L, Simons K. Lipid rafts and membrane dynamics. J. Cell Sci. 2005; 118:1099-1102.

Zeyda M, Stulnig TM. Lipid Rafts & Co.: an integrated model of membrane organization in T cell activation. Prog. Lipid Res. 2006; 45:187-202.

 

See Also

Membrane Assembly and Stability

Signal Cascades, Protein Interaction Networks in

Lipid Rafts

Synthetic Lipids to Study Biological Function