Visualization of Annexin I Binding to Calcium‐Induced Phosphatidylserine Domains

Annexins are a family of structurally related eukaryotic proteins that reversibly bind membranes containing anionic phospholipids in a calcium-dependent manner. More than 160 different isoforms have been found in many organisms ranging from mammals to molds. The protein family is defined by its characteristic structure comprising a conserved core made up of four or eight domains of a 70-amino-acid sequence forming five a helices and a variable N-terminal region varying in length and amino acid sequence. The core domains harbor multiple calcium-binding sites, which are all located on the convex side of the molecule. X-ray crystallographic analyses and mutagenesis studies have shown that the convex site is responsible for initial membrane binding. Calcium ions bound to these sites act as bridges connecting the protein with anionic lipid headgroups. The N-terminal region is thought to be involved in the regulation of different functions of annexins. Although exact physiological functions of annexins have not been identified yet, it has been shown that they participate in a variety of in vitro activities. In particular, some annexins such as annexins I, II, IV and VII can promote membrane aggregation and may thus be involved in cellular endoand exocytotic pathways. It was shown that annexin I, the protein of interest in our study, is capable of aggregating and even fusing membrane vesicles. However, the mechanism of membrane aggregation is still discussed controversially. One model postulates that membrane-bound annexin I molecules form axial dimers prior to interacting with a second membrane, while another model hypothesizes that monomeric bound annexin I interacts with the second membrane. We utilized scanning force microscopy (SFM) on solidsupported Langmuir ± Blodgett (LB) bilayers composed of 1,2dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and 1,2-dipalmitoyl-sn-glycero-3-phosphoserine (DPPS) immobilized on mica (serving as an atomically flat substrate) to directly visualize annexin I binding with high lateral and vertical resolution, thus enabling us to distinguish between the two different models. With this technique height differences in the aÊngström region as well as morphological changes of the membrane structure and domain formation can be observed in a physiological environment. Topographic images of an LB bilayer composed of DPPC as the first leaflet and DPPC/DPPS (4:1) representing the uppermost layer pointing to the aqueous phase are mostly featureless, with some defects occurring as dark spots in the SFM images. Addition of a 1 mM annexin I solution in 50 mM Tris (pH 7.4), 1 mM CaCl2 to the bilayer results in the appearance of circular domains with sizes of 3 ± 10 mm (Figures 1 a and b) that are attributed to specifically adsorbed annexin I. These protein domains occupy (35 3) % of the overall area, exhibit an average height of (3.2 0.3) nm as obtained from a height analysis (Figure 1 c), and are stable in a calcium-containing buffer for several hours. The thickness of the annexin layer compares well with the molecular dimension of annexin I as determined by X-ray crystallography, indicating that the protein binds in a monomeric fashion. 17] Recently, Bitto et al. , employing X-ray specular reflectivity measurements, also found that annexin I binds as a monomer or monolayer to 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoserine (POPS)/1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE)/1-palmitoly-2-oleoyl-sn-glycero-3-phosphocholine (POPC) (2:5:2) monolayers at the air ± water interface. They determined a protein thickness of (3.1 0.2) nm. Remarkably, lateral movement of the protein domains was not observed in the Ca2-containing buffer within three hours indicating that the lateral mobility of the lipids is low, as is expected for gel-phase lipids at room temperature. Crystallization of the protein, however, did not occur as it was reported for annexin V on a DOPC/DOPS (4:1) membrane immobilized on mica. 20] In [23] J. D. Thompson, D. G. Higgins, T. J. Gibson, Nucleic Acids Res. 1994, 22, 4673 ± 4680. [24] F. Jeanmougin, J. D. Thompson, M. Gouy, D. G. Higgins, T. J. Gibson, Trends Biochem. Sci. 1998, 23, 403 ± 405. [25] D. A. Berthold, M. E. Andersson, P. Nordlund, Biochem. Biophys. Acta 2000, 1460, 241 ± 254. [26] N. Guex, M. C. Peitsch, Electrophoresis 1997, 18, 2714 ± 2723. [27] WebLab ViewerLite, Molecular Simulations, Inc. , Princeton, 1998.