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Capturing suboptical dynamic structures in lipid bilayer patches formed from free-standing giant unilamellar vesicles

Nature Protocols volume 12, pages 1563–1575 (2017)Cite this article

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Abstract

There is accumulating evidence that the small-scale lateral organization of biological membranes has a crucial role in signaling and trafficking in cells. However, it has been difficult to characterize these features with existing methods for preparing and analyzing freestanding membranes, because the dynamics occurs below the optical resolution possible with these protocols. We have developed a protocol that permits the imaging of lipid nanodomains and lateral protein organization in membranes of giant unilamellar vesicles (GUVs). Freestanding GUVs are transferred onto a mica support, and after treatment with magnesium chloride, they collapse to form planar lipid bilayer (PLB) patches. Rapid GUV collapse onto the mica preserves the lateral organization of freestanding membranes and thus makes it possible to image 'snapshots' of GUVs up to nanometer resolution by high-resolution microscopy. The method has been applied to classical lipid raft mixtures in which suboptical domain fluctuations have been imaged in both the liquid-ordered and liquid-disordered membrane phases. High-resolution scanning by atomic force microscopy (AFM) of membranes composed of binary and ternary lipid mixtures reconstituted with Na+/K+-ATPase (NKA) has revealed the spatial distribution and orientations of individual proteins, as well as details of membrane lateral structure. Immunolabeling followed by confocal microscopy can also provide information about the spatial distribution of proteins. The protocol opens up a new avenue for quantitative biophysical studies of suboptical dynamic structures in biomembranes, which are local and short-lived. Preparation of GUVs, PLB patches and their imaging takes <24 h.

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Figure 1: Orientation of NKA in a vesicle.
Figure 2: The fluorescence intensity of the membrane probes RhPE and naphthopyrene on the surface of GUVs used to quantify the area fraction of the lo (by naphthopyrene) and the ld (by RhPE) domains for a batch of vesicles.
Figure 3: Epifluorescence images of GUVs (numbered 1–4) before and after collapse.
Figure 4: Epifluorescence and AFM images of a planar bilayer patch of GUVs displaying macroscopic and suboptical lo–ld domains.
Figure 5: AFM topography images and immunolabeling of a PLB patch, showing the spatial distribution and orientation of NKA in the membranes of binary and ternary GUVs.

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References

  1. Simons, K. & Ikonen, E. Functional rafts in cell membranes. Nature 387, 569–572 (1997).

    Article  CAS  Google Scholar 

  2. Hanzel-Bayer, M.F. & Hancock, J.F. Lipid rafts and membrane traffic. FEBS Lett. 581, 2098–2104 (2007).

    Article  Google Scholar 

  3. Brown, D.A. & London, E. Structure and function of sphingolipid- and cholesterol-rich membrane rafts. J. Biol. Chem. 275, 17221–17224 (2000).

    Article  CAS  Google Scholar 

  4. Eggeling, C. et al. Direct observation of the nanoscale dynamics of membrane lipids in a living cell. Nature 457, 1159–1162 (2009).

    Article  CAS  Google Scholar 

  5. Ringemann, C. et al. Exploring single-molecule dynamics with fluorescence nanoscopy. New J. Phys. 11, 103054 (2009).

    Article  Google Scholar 

  6. Ipsen, J.H., Jorgensen, K. & Mouritsen, O.G. Density fluctuations in saturated phospholipid bilayers increase as the acyl-chain length decreases. Biophys. J. 58, 1099–1107 (1990).

    Article  CAS  Google Scholar 

  7. Gil, T. et al. Theoretical analysis of protein organization in lipid membranes. Biochim. Biophys. Acta 1376, 245 (1998).

    Article  CAS  Google Scholar 

  8. Hønger, T. et al. Anomalous swelling of multilamellar lipid bilayers in the transition region by renormalization of curvature elasticity. Phys. Rev. Lett. 72, 3911–3914 (1994).

    Article  Google Scholar 

  9. Cruzeiro-Hansson, L., Ipsen, J.H. & Mouritsen, O.G. Intrinsic molecules in lipid-membranes change the lipid-domain interfacial area: cholesterol at domain interfaces. Biochim. Biophys. Acta 979, 166–176 (1989).

    Article  CAS  Google Scholar 

  10. Vist, M. & Davis, J. Phase-equilibria of cholesterol dipalmitoylphosphatidylcholine mixtures—H2 nuclear magnetic-resonance and differential scanning calorimetry. Biochemistry 29, 451–464 (1990).

    Article  CAS  Google Scholar 

  11. Almeida, P., Vaz, W. & Thompson, T. Lateral diffusion in the liquid-phases of dimyristoylphosphatidylcholine cholesterol lipid bilayers—a free-volume analysis. Biochemistry 31, 6739–6747 (1992).

    Article  CAS  Google Scholar 

  12. Gowrishankar, K. et al. Active remodeling of cortical actin regulates spatiotemporal organization of cell surface molecules. Cell 149, 1353–1367 (2012).

    Article  CAS  Google Scholar 

  13. Dietrich, C., Yang, B., Fujiwara, T., Kusumi, A. & Jacobson, K. Relationship of lipid rafts to transient confinement zones detected by single particle tracking. Biophys. J. 82, 274–284 (2002).

    Article  CAS  Google Scholar 

  14. Tokumasu, F., Jin, A., Feigenson, G. & Dvorak, J. Nanoscopic lipid domain dynamics revealed by atomic force microscopy. Biophys. J. 84, 2609–2618 (2003).

    Article  CAS  Google Scholar 

  15. Dietrich, C. et al. Lipid rafts reconstituted in model membranes. Biophys. J. 80, 1417–1428 (2001).

    Article  CAS  Google Scholar 

  16. Tamm, L. & McConnell, H. Supported phospholipid-bilayers. Biophys. J. 47, 105–113 (1985).

    Article  CAS  Google Scholar 

  17. Milhiet, P., Giocondi, M. & Grimellec, C. AFM imaging of lipid domains in model membranes. Sci. World J. 17, 59 (2003).

    Article  Google Scholar 

  18. Alessandrini, A. & Facci, P. Phase transitions in supported lipid bilayers studied by AFM. Soft Matter 10, 7145–7164 (2014).

    Article  CAS  Google Scholar 

  19. Heinemann, F. & Schwille, P. Preparation of micrometer-sized free-standing membranes. Chem. Phys. Chem. 12, 2568–2571 (2011).

    Article  CAS  Google Scholar 

  20. Papahadjopoulos-Sternberg, B. Freeze-fracture electron microscopy on domains in lipid mono- and bilayer on nano-resolution scale. Methods Mol. Biol. 606, 333–349 (2010).

    Article  CAS  Google Scholar 

  21. Bhatia, T., Husen, P., Ipsen, J.H., Bagatolli, L.A. & Simonsen, A.C. Fluid domain patterns in free-standing membranes captured on a solid support. Biochim. Biophys. Acta 1838, 2503–2510 (2014).

    Article  CAS  Google Scholar 

  22. Bhatia, T. et al. Spatial distribution and activity of Na+K+-ATPase in lipid bilayer membranes with phase boundaries. Biochim. Biophys. Acta 1858, 1390–1399 (2016).

    Article  CAS  Google Scholar 

  23. Jass, J., Tjärnhage, T. & Puu, G. From liposomes to supported planar bilayer structures on hydrophilic and hydrophobic surfaces: an atomic force microscopy study. Biophys. J. 79, 3153–3163 (2000).

    Article  CAS  Google Scholar 

  24. Horn, R.G. Direct measurement of the force between two lipid bilayers and observation of their fusion. Biochim. Biophys. Acta 778, 224–228 (1984).

    Article  CAS  Google Scholar 

  25. Richter, R.P. & Brisson, A.R. Following the formation of supported lipid bilayers on mica: a study combining AFM, QCM-D, and ellipsometry. Biophys. J. 88, 3422–3433 (2005).

    Article  CAS  Google Scholar 

  26. Hamai, C., Cremer, P.S. & Musser, S.M. Single giant vesicle rupture events reveal multiple mechanisms of glass-supported bilayer formation. Biophys. J. 92, 1988–1999 (2007).

    Article  CAS  Google Scholar 

  27. Xie, Z. Molecular mechanisms of Na+,K+-ATPase mediated signal transduction. Ann. N. Y. Acad. Sci. 986, 497–503 (2003).

    Article  CAS  Google Scholar 

  28. Skou, J. The influence of some cations on an adenosine triphosphatase from peripheral nerves. Biochim. Biophys. Acta 23, 394–401 (1957).

    Article  CAS  Google Scholar 

  29. Cornelius, F. In Biomimetic Membranes for Sensor and Separation Applications, Biological and Medical Physics, Biomedical Engineering (ed. Hèlix-Nielsen, C.), 6, 113–135 (Springer, 2012).

    Google Scholar 

  30. Cornelius, F., Habeck, M., Kanai, R., Toyoshima, C. & Karlish, S.J.D. General and specific lipid-protein interactions in NaK-ATPase. Biochim. Biophys. Acta 1848, 1729–1743 (2015).

    Article  CAS  Google Scholar 

  31. Cornelius, F. Functional reconstitution of the sodium pump kinetics of exchange reactions performed by reconstituted NaK-ATPase. Biochim. Biophys. Acta 1071, 19–66 (1991).

    Article  CAS  Google Scholar 

  32. Kanai, R., Ogawa, H., Vilsen, B., Cornelius, F. & Toyoshima, C. Crystal structure of a Na+-bound Na+K+-ATPase preceding the E1P state. Nature 502, 201–206 (2013).

    Article  CAS  Google Scholar 

  33. Shinoda, T., Ogawa, H., Cornelius, F. & Toyoshima, C. Crystal structure of the sodium-potassium pump at 2.4 Å resolution. Nature 459, 446–450 (2009).

    Article  CAS  Google Scholar 

  34. Bhatia, T. et al. Preparing giant unilamellar vesicles of complex lipid mixtures on demand: mixing small unilamellar vesicles of compositionally heterogeneous mixtures. Biochim. Biophys. Acta 1848, 3175–3180 (2015).

    Article  CAS  Google Scholar 

  35. Bhatia, T., Cornelius, F., Mouritsen, O.G. & Ipsen, J.H. Reconstitution of transmembrane protein Na+,K+-ATPase in giant unilamellar vesicles of lipid mixtures involving PSM, DOPC, DPPC and cholesterol at physiological buffer and temperature conditions. Protoc. exch. http://dx.doi.org/10.1038/protex.2016.010 (2016).

  36. Husen, P., Arriaga, L.R., Monroy, F., Ipsen, J.H. & Bagatolli, L.A. Morphometric image analysis of giant vesicles, a new tool for quantitative thermodynamics studies of phase separation in lipid membranes. Biophys. J. 103, 2304–2310 (2012).

    Article  CAS  Google Scholar 

  37. Husen, P., Arriaga, L.R., Monroy, F., Ipsen, J.H. & Bagatolli, L.A. A method for analysis of lipid vesicle domain structure from confocal image data. Eur. Biophys. J. 41, 161–175 (2012).

    Article  CAS  Google Scholar 

  38. Fidorra, M., Garcia, A., Ipsen, J.H., Härtel, S. & Bagatolli, L.A. Lipid domains in giant unilamellar vesicles and their correspondence with equilibrium thermodynamic phases: a quantitative fluorescence microscopy imaging approach. Biochim. Biophys. Acta 1788, 2142–2149 (2009).

    Article  CAS  Google Scholar 

  39. Bhatia, T., Cornelius, F. & Ipsen, J.H. Exploring the Raft-hypothesis by probing planar bilayer patches of free-standing giant vesicles at nanoscale resolution, with and without Na,K-ATPase. Biochim. Biophys. Acta 1858, 3041–3049 (2016).

    Article  CAS  Google Scholar 

  40. Cornelius, F. Incorporation of C12E8-solubilized Na+,K+-ATPase into liposomes, determination of sidedness and orientation. Methods Enzymol. 156, 156–167 (1988).

    Article  CAS  Google Scholar 

  41. Cornelius, F. & Moller, J.V. In Handbook of Non-medical Applications of Liposomes Vol. 2 (eds. Lasic, D.D. & Barenholz, Y.) 219–243 (CRC Press, 1995).

    Google Scholar 

  42. Cornelius, F. Cholesterol dependent interaction of polyunsaturated phospholipids with NaK-ATPase. Biochemistry 47, 1652–1658 (2008).

    Article  CAS  Google Scholar 

  43. Powalska, E. et al. Fluorescence spectroscopic studies of pressure effects on NaK-ATPase reconstituted into phospholipid bilayers and model raft mixtures. Biochemistry 46, 1672–1683 (2007).

    Article  CAS  Google Scholar 

  44. Honerkamp-Smith, A.R. et al. Line tensions, correlation lengths, and critical exponents in lipid membranes near critical points. Biophys. J. 95, 236–246 (2008).

    Article  CAS  Google Scholar 

  45. Palmieri, B. et al. Line active molecules promote inhomogeneous structures in membranes: theory, simulations and experiments. Adv. Colloid Interface Sci. 208, 58–65 (2014).

    Article  CAS  Google Scholar 

  46. Safran, S. Micelles Membranes Microemulsions and Monolayers. Springer, 1994.

  47. Charras, G.T., Coughlin, M., Mitchison, T.J. & Mahadevan, L. Life and times of a cellular bleb. Biophys. J. 94, 1836–1853 (2008).

    Article  CAS  Google Scholar 

  48. Møller, J.V. et al. Probing of the membrane topology of sarcoplasmic reticulum Ca2+-ATPase with sequence-specific antibodies. J. Biol. Chem. 272, 29015–29032 (1997).

    Article  Google Scholar 

  49. Muller, D.J. & Engel, A. Atomic force microscopy and spectroscopy of native membrane proteins. Nat. Protoc. 2, 2191–2197 (2007).

    Article  Google Scholar 

  50. Veatch, S.L., Polozov, I.V., Gawrisch, K. & Keller, S.L. Liquid domains in vesicles investigated by NMR and fluorescence microscopy. Biophys. J. 86, 2910–2922 (2004).

    Article  CAS  Google Scholar 

  51. Apell, H.J. & Bersch, B. Oxonol VI as an optical indicator for membrane potentials in lipid vesicles. Biochim. Biophys. Acta 903, 480–494 (1987).

    Article  CAS  Google Scholar 

  52. Peterson, G.L. A simplification of the protein assay method of Lowry et al. which is more generally applicable. Anal. Biochem. 83, 346–356 (1977).

    Article  CAS  Google Scholar 

  53. Lowry, O.H. Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265–275 (1951).

    CAS  PubMed  Google Scholar 

  54. Baginski, E.S., Foa, P.P. & Zak, B. Microdetermination of inorganic phosphate, phospholipids, and total phosphate in biologic materials. Clin. Chem. 13, 326–332 (1967).

    CAS  PubMed  Google Scholar 

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Acknowledgements

We thank DAMBIC (Danish Molecular Bioimaging Center) for access to equipment. This work was supported by the Danish Council for Independent Research–Natural Sciences (FNU; grant 95-305-23443 to J.H.I. and F.C.), the Human Frontier Science Program (HFSP; grant 95-305-73458 to J.H.I.) and the Novo Nordisk Foundation (grant DFF-418300011 to F.C.), T.B. acknowledges O.G. Mouritsen (SDU), A.C. Simonsen (SDU), P.L. Hansen (SDU), L.A. Bagatolli (SDU), J. Brewer (SDU), L. Duelund (SDU), B. Franchi (AU) and H. Kidmose (AU) for useful discussions.

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Authors and Affiliations

  1. MEMPHYS Center for Biomembrane Physics and Department of Physics, Chemistry, and Pharmacy, University of Southern Denmark, Odense, Denmark

    Tripta Bhatia & John H Ipsen

  2. Max Planck Institute of Colloids and Interfaces, Potsdam, Germany

    Tripta Bhatia

  3. Department of Biomedicine, Aarhus University, Aarhus, Denmark

    Flemming Cornelius

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Contributions

T.B. performed the research and analyzed the data; designed the vesicle collapse method and developed it through many useful discussions with J.H.I.; and developed the SUV-mixing method together with J.H.I. F.C. prepared and characterized proteoliposomes for their functional and structural properties; prepared the primary antibodies for NKA; and discussed the protein-reconstitution protocol, antibody-labeling experiments and NKA data with T.B. J.H.I. planned the experiments with T.B. and contributed to data interpretation. T.B. and J.H.I. wrote the paper.

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Correspondence to Tripta Bhatia.

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Supplementary information

Time sequences showing GUV collapse after the addition of MgCl2 salt

The starting frame (t = 0 s) shows GUVs of ternary lipid mixture DOPC-DPPC-chol (30-50-20)% sitting on mica substrate displaying macroscopic ld (bright) and lo (dark) domains. There is no MgCl2 salt present in the fluid chamber. GUV collapse starts at t ∼ 3.820 s after the addition of MgCl2 salt, and planar lipid bilayer patches are formed that retain macroscopic ld (bright) and lo (dark) domains. GUVs contain the membrane probe RhPE, which preferentially partitions into the ld phase and is imaged as a bright region in GUVs and patches. GUVs were imaged in an epifluorescence microscope equipped with an EMCCD camera. Scale bar, 10 μm. Reproduced with permission from ref. 21. (AVI 1577 kb)

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Bhatia, T., Cornelius, F. & Ipsen, J. Capturing suboptical dynamic structures in lipid bilayer patches formed from free-standing giant unilamellar vesicles. Nat Protoc 12, 1563–1575 (2017). https://doi.org/10.1038/nprot.2017.047

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