Flavel, Benjamin Scott

Miners, John Oliver

Nussio, Matthew Robert

Shapter, Joseph George

Sykes, Matthew James

Voelcker, Nicolas Hans

Flinders Academic Commons

High Resolution Chemical Mapping of Biomimetic Membranes by Force Volume Imaging Matthew R. Nussio*, Nicolas H. Voelcker*, Matthew J. Sykes, Ben S. Flavel*, John O. Miners, and Shapter* *School of Chemistry, Physics and Earth Sciences Department of Clinical Pharmacology Flinders University, Sturt Road, Bedford Park, Adelaide, SA 5001 Email: joe.shapter@flinders.edu.au    Abstract The analysis of membrane domain segregation and phase separation is of significant interest for the understanding of cell membranes. In this paper, we report a new approach to atomic force microscopy (AFM) of artificial membranes, using both functionalised AFM tips and Force Volume AFM imaging for the analysis of membrane phase separation. Simultaneous topology and mapping of interaction forces of binary component phospholipid bilayer membranes was performed yielding novel insights into membrane structure at unprecedented resolution. Keywords-phospholipids; microscopy; force-volume I. INTRODUCTION  The lack of reliable chemical analysis adapted to the challenging task of analysing cell membrane lipid composition is a key problem in membrane research. This problem is exacerbated by the fact that each leaflet of the cells bilayer comprises a different lipid composition. Techniques that allow the in situ probing or mapping of these membrane components are scarce [1]. The time dynamics and variation of phase and chemical composition in this system demand fast, high resolution techniques. In-fluid AFM combined with spectroscopy (so called Force Volume (FV) mode) can satisfy these demands. However, FV imaging has been limited to producing elasticity maps of heterogeneous materials including biominerals [2], carbon nanotubes [3], polymer composites [4-6] and living cells [7, 8]. Here, we present a new approach for the analysis of model membranes based on FV imaging using AFM tips with defined surface chemistry which will allow unprecedented insights into bilayer membrane organization and composition. This will provide invaluable new insights into lipid dynamics at levels not previously possible. II. EXPERIMENTAL  A. Vesicle Preparation Multilamellar vesicles (MLVs) were prepared by first dissolving aliquots of dipalmitoyl phosphatidylcholine (DPPC) and/or dioleoyl phosphatidylserine (DOPS) (Avanti Polar Lipids, Birmingham, AL, USA) in chloroform, followed by evaporation of the solvent under nitrogen. Lipid samples were further dried under vacuum for 3 h prior to being suspended in 10 mM HEPES buffer, 150 mM NaCl,  4mM CaCl2. The final concentration of lipids was 0.5 mM. Samples were left to hydrate overnight, followed by sonication for 30 min. During sonication, periodic vortex mixing was carried out prior to vesicle extrusion (Avanti Mini-Extruder, Avanti Polar Lipids, Birmingham, AL, USA) 20 times through a polycarbonate membrane filter of defined pore diameter, typically 100 nm. Extrusion was performed at temperatures above the transition temperature (TM) of the component phospholipids, since gel-state lipids are difficult to extrude at lower temperatures [9]. Resultant small unilamellar vesicles (SUVs) had a monodisperse size distribution as measured by dynamic light scattering (HPPS, Malvern Instruments) and were used for all further experiments.  B. Atomic Force Microscopy The visualization of SPBs was performed using a commercial AFM (Nanoscope IV, Digital Instruments, Santa Barbara, CA). All images were obtained by means of in situ tapping mode using triangular Si3N4 cantilevers (Digital Instruments) with a spring constant of 0.15 N m-1 operating at the cantilever resonant frequency. Formation of SPBs was achieved by depositing 100L of 100nm SUV solution (10mM HEPES, 150mM NaCl, 4mM CaCl2, pH 7) to a freshly cleaved mica surface. Prepared surfaces were then incubated for 4hrs at 4oC [10]. Prior to imaging, surfaces were rinsed with 10mM HEPES, 150mM NaCl, pH 7. C. Chemical Modification of AFM Probes. Commercial triangular Si3N4 cantilevers were cleaned in a water plasma prior to further modification. Following plasma cleaning, tips were modified using silane deposition from solution [11]. Modification of plasma treated tips with 3-aminopropyltriethoxysilane (APTES) (0.5% v/v in hexadecane) was performed by deposition overnight (16h) at room temperature followed by rinsing in chloroform and drying under a gentle stream of N2.   Joseph G. 1-4244-1504-7/08/$25.00 © 2008 IEEE ICONN 2008121D. Force Spectroscopy and Force Volume Force plots were acquired using either triangular Si3N4 cantilevers or APTES modified Si3N4 cantilevers with a nominal spring constant of 0.15 N m-1. A total of approximately 200 force curves over more than five positions were obtained for each surface analysed. Force volume imaging collects force curves at each x,y pixel of the image by measuring cantilever deflection. Whilst a height image is also produced by this technique, its strength is that the information contained in the 3D data set can be decoupled from the topographic information.  All force spectroscopy and force volume measurements were performed in 10mM HEPES, 1mM NaCl pH 7. In this work, we focus on the force data obtained for the retract cycle. III. RESULTS & DISCUSSION  Elucidating the role of the fine structure and composition of cellular membranes has been of significant interest in the past few decades. Exactly how membrane domains influence the transduction of cellular signals is under constant investigation. The involvement of membranes in diverse, critical cellular functions justifies the existence of a spectrum of constituent phospholipids, and the ability to understand their spatial distribution and physical properties is key to ultimately understanding physiological multi-component lipid membranes. In addressing the current lack of understanding of membrane organisation and its function, the characterisation of the spatial organisation o f membrane lipids with nanoscale feature resolution has been performed for binary mixtures of phosphatidylcholine and phosphatidylserine.  For the current study, a 1:1 binary mixture of DPPC and DOPS was prepared. As expected, supported phospholipid bilayers from this lipid mixture exhibited phase separation such that the DPPC gel domains were typically ~1nm higher than the neighbouring DOPS liquid phase domains (Figure 1) [10].   Figure 1.  AFM topographic image ( 1.5 x 1.5 m2; Z-scale: 5 nm, 512 lines x 512 lines) of a 50:50 DPPC:DOPS supported phospholipid bilayer. DPPC domains extended 1.18±0.03 nm above surrounding DOPS domains.  Zooming in to image a smaller region of the bilayer, Figure 2A shows a low resolution topographic image. Figure 2B is the corresponding FV image obtained using a chemically-modified tip where the adhesion force is mapped as a function of lateral position.                    Figure 2.  (A) Topographic height image (32 lines x 32 lines) and (B) FV adhesion map (322 forces curves) of a 50:50 DPPC:DOPS supported phospholipid bilayer. Lighter coloured regions in the FV adhesion map correspond to greater force associated with DOPS phospholipids. Since the tip modification affords two very different adhesion forces for each component phospholipid (Figure 3), the identification of lipids in the FV image is facilitated.            Figure 3.  Representative retraction force curves for (A) DOPS, and (B) DPPC obtained from an APTES modified probe. Inset: molecular structures of each phospholipid. 1nN 70nm DOPS DPPC AB BNanodomains AB122The difference in response towards tip surface chemistry can be attributed to the difference in functionality of the head groups in the two lipids (Figure 3). The tertiary amines of DPPC limit hydrogen bonding to the modified AFM tip and hence little adhesion is observed (dark areas in Figure 2B). However, in the case of DOPS, the lipids carboxylic acid readily hydrogen bond to the amine-terminated tip, giving rise to a large adhesive force (light areas in Figure 2A). In the upper part of the FV image (Figure. 2B), nanoscale DOPS domains within a larger DPPC domain are apparent.  These nanodomains are on the order of 100 nm in lateral dimensions and cannot be resolved at this level of detail by any other technique. Measuring the mobility of dye-labelled lipid molecules using FCS, DPPC domains within a DOPC matrix have previously identified locations where the mobility of the lipid probes were almost the same as the liquid phase DOPC regions [12]. It was inferred that the biphasic mobility of the observed nanoscale DPPC domains may have been due to DPPC packing heterogeneities or DOPC clusters surrounding lipid fluorescent probes. The results reported in this paper suggest that this measured change may have been due to nanodomains absorbed into larger microdomains. A recent study utilising NanoSIMS has also revealed the existence of a fluid phase subdomain within a gel phase domain, which was assumed to represent trapping of the fluid-phase subdomain upon expansion of the gel-phase domain [13].    With reference to the FV image (Figure 2B), the adhesion forces can be mapped along a line moving horizontally from left to right (Figure 4). The change of adhesion force when moving the tip across the interface from one lipid domain to the other is conspicuous.  Figure 4.  Adhesion force as a function of position for several lines in FV adhesion map (Figure 2B). Several lines demonstrate a gradual decline in adhesion at the interface, implying that intermixing between DOPS and DPPC bilayer domains take place. For lines in the upper part of the image, there is a pronounced step in adhesion from one region of lipid to the other. However, in the lower part of the image, the change in adhesion across the step is gradual and several values between the two extremes of the two pure lipids are observed. This result indicates that in fact the lipid domain boundary in this region is not as well defined, consistent with significant intermixing of the two lipids. This intermixing occurs over a region of about 40 nm and may influence bilayer properties significantly. Studies utili sing FCS have hinted at heterogeneities of lipid packing at the domain boundary of binary mixtures of DOPC and DPPC [12]. The herein described, to date unknown, lipid intermixing phenomenon may also play an important role in the formation and movement of lipid rafts or in the formation and functional behaviour of protein assemblies in the complex and phase separated structures of physiological membranes. Association of proteins into distinct lipid microdomains is governed by either chemical or physical properties of the lipid microdomain environment. For example, it is understood that glycosylphosphatidyl (GPI)-anchored proteins partition enrich lipid rafts, while many transmembrane proteins are located in nonraft phases [14]. Proteins can also be confined to the area of the microdomain if the activation energy of passing the domain barrier is larger than the kinetic energy of the protein itself [15]. Hypothetically, the intermixing boundary observed in the current study could potentially provide a mechanism for proteins in different lipid environments to interact. With the aid of a specialised AFM imaging technique known as FV imaging, we have imaged the topology and mapped the interaction forces of binary component phospholipid bilayer membranes at a lateral resolution of 20nm, a more than twofold improvement over the current state-of-the-art. A binary mixture of phosphatidylcholine and phosphatidylserine was utilised in these experiments but the continued development of this technique has the potential to examine multiple headgroup functionalities, eventually leading to the compositional analysis of physiologically relevant biological membranes.  Development of this technique to monitor physiological relevant membranes will have potential applications in elucidating the nature and role of cellular membranes, in particular their domains or rafts, and how their dynamic structure and molecular associations affect cell membrane signalling or biological functions.  ACKNOWLEDGMENT The authors wish to thank Dr. Mickey Huson and co-workers from CSIRO (Textile and Fibre Technology) for providing and offering assistance with the software Force Volume.  REFERENCES [1] G.W. Feigenson, Phase behavior of lipid mixtures Nature Chemical Biology, 2006. 2: p. 560-563.  [2] A.N. Parbhu, W.G. Bryson, R. Lal, Disulfide Bonds in the Outer Layer of Keratin Fibers Confer Higher Mechanical Rigidity: Correlative Nano-Indentation and Eleasticity Measureme nt with an AFM Bio chemistry, 1999. 38: p. 11755-11761. [3] M.A. Poggi, P.T. Lillehei, L.A. 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Lal, Physiological Role of Gap-junctional Hemichannels: Extracellular Calcium-dependent Isosmotic Volume Regulation, The Journal of Cell Biology, 2000. 148: p. 1053-1074. [8] C.V. Gopal Reddy, K. Malinowska, N. Menhart, R. Wang, Identification of TrkA on living PC12 cells by atomic force microscopy, Biochimica et Biophysica Acta, 2004. 1667: p. 15-25. [9] R.C. MacDonald, R.I. MacDonals, B.P. Menco, K. Takeshita, N.K. Subbarao, L.R. Hu, Small -volume extrusion apparatus for preparation of large unilamellar vesicles Biochimica et Biophysica Acta, 1991. 1061: p. 297-303. [10] I. Reviakine, A. Simon, A. Brisson, Effect of Ca2+ on the Morphology of Mixed DPPC-DOPS Supported Phospholipid Bilayers Langmuir, 2000. 16: p. 1473-1477. [11] D. Losic, K. Short, J.G. Shapter, J.J. Gooding, Atomic Force Microscopy Imaging of Glucose Oxidase using Chemically Modified Tips, Atomic Force Microscopy Imaging of Glucose Oxidase using Chemically Modified Tips, 2003. 56: p. 1039-1043. [12] A.R. Burns, D.J. Frankel, T. Buranda, Local Mobility in Lipid Domains of Supported Bilayers Characterized by Atomic Force Microscopy and Fluorescence Correlation Spectroscopy Biophysical Journal, 2005. 89: p. 1081 -1093. [13] M.L. Kraft, P.K. Weber, M.L. Longo, I.D. Hutcheon, S.G. Boxer, Phase Separation of Lipid Membranes Analyzed with High-Resolution Secondary Ion Mass Spectrometry Science, 2006. 313: p. 1948-1951. [14] J.J. Sieber, K.I. Willig, C. Kutzner, C. Gerding-Reimers, B. Harke, G. Donnert, B. Rammner, C. Eggeling, S.H. Hell, H. Grubmuller, T. Lang, Anatamy and Dynamics of a Supramolecular Membrane Protein Cluster Science, 2007. 317: p. 1072-1076. [15] G. Vereb, J. Szollosi, J. Matko, P. Nagy, T. Farkas, L. Vigh, L. Matyus, T.A. Waldmann, Dynamic, yet structured: The cell membrane three decades after the Singer-Nicolson model Proceedings of the National Academy of Sciences of the Unites States of America, 2003. 100: p. 8053-8058.  124

High resolution chemical mapping of biomimetic
            membranes by force volume imaging

Matthew R. Nussio

Nicolas H. Voelcker

Matthew J. Sykes

Ben S. Flavel

John O. Miners

Joseph G. Shapter

Crossref

High resolution chemical mapping of biomimetic membranes by force volume imaging

Nussio, Matthew R.

Voelcker, Nicolas H.

Sykes, Matthew J.

Flavel, Ben S.

Miners, John O.

Shapter, Joseph G.

University of Queensland eSpace

http://dspace.flinders.edu.au/xmlui/bitstream/handle/2328/9057/2006009183.pdf?sequence=1

High resolution chemical mapping of biomimetic membranes by force volume imaging

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