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* Wellcome Trust Centre for Cell-Matrix Research, School of Biological Sciences, and School of Medicine, University of Manchester, Manchester, M13 9PT, United Kingdom
Correspondence: Address reprint requests to Michael J. Sherratt, Tel.: +44-161-275-7054; Fax: +44-161-275-5082; E-mail: michael.sherratt{at}man.ac.uk.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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) and that in physiological environments many processes, such as those occurring at the cell surface, are dependent on interfacial phenomena (Kowalewski and Holtzman, 1999). These substrate interactions have the potential to induce conformational changes in adsorbed proteins (Haynes and Norde, 1995), which can affect biological function (Grinnell and Feld, 1981; Garcia et al., 1999; Keselowsky et al., 2003).
Physisorption of proteins to substrates is driven by van der Waals forces, the electrostatic double-layer (EDL) force and the hydrophobic effect (Muller et al., 1997). The van der Waals force can be either attractive or repulsive and the magnitude of the force is dependent on the geometry of the interacting features (Leckband, 2000). The EDL force is dependent on the concentration and valency of charged solutes and the surface charge density of both substrate and protein (Muller et al., 1997). Hydrophobic effects are thought to provide the dominant driving force for the folding of globular proteins in solution where hydrophobic side groups tend to be buried in the interior and hydrophilic side groups are usually located on the periphery of the protein (Haynes and Norde, 1995). The adsorption of proteins to solid surfaces (particularly hydrophobic surfaces) may involve the formation of a water-free contact layer where the protein can unfold without solvating all of its hydrophobic residues (Haynes and Norde, 1995). Substrate hydrophobicity/hydrophilicity has been shown to affect the morphology and/or biological function of adsorbed synthetic and biological macromolecules. The height of adsorbed synthetic ethylenediamine core poly(amidoamine) dendrimers approached the theoretical sphere diameter on hydrophobic but not hydrophilic substrates (Betley et al., 2001). On hydrophilic mica, Alzheimer's ß-amyloid peptide formed particulate, pseudomicellar aggregates, but on hydrophobic graphite the same protein organized into uniform, elongated sheets (Kowalewski and Holtzman, 1999).
Substrate dependent morphologies and functions have also been demonstrated for some mammalian extracellular matrix (ECM) proteins. Recombinant human elastin peptides coacervate in heated solutions to form fibrillar aggregates (Bellingham et al., 2001). In an atomic force microscopy (AFM) study these elastin peptides adsorbed to a hydrophilic surface as discrete, rounded aggregates but on a hydrophobic surface the peptides self-assembled into an energetically favorable hexagonally closed packed fibril arrangement (Yang et al., 2002). Fibronectin is an important modular ECM protein, which binds to integrins, collagen, heparan sulfate, hyaluronic acid, and itself. The binding of integrins to surface-bound fibronectin is dependent on the hydrophilicity/hydrophobicity of the surface. Substrate-specific conformational changes have been inferred from solid-phase cell binding assays (Grinnell and Feld, 1981; Garcia et al., 1999) and demonstrated for adsorbed fibronectin by AFM (Bergkvist et al., 2003).
Although substrate-dependent morphologies have been observed for some isolated proteins, these effects have not been described for the functional supramolecular assemblies formed by most ECM components. Fibrillin microfibrils and type-VI collagen microfibrils are two biochemically distinct ECM supramolecular assemblies, which interact with cells and other ECM components in vitro and in vivo (Kielty et al., 2002a; Baldock et al., 2003). The interest in fibrillin microfibril biology has been driven by the need to understand the fundamental mechanisms of tissue elasticity and the basis of genotype-to-phenotype correlations in heritable connective tissue diseases such as Marfan syndrome, which are caused by mutations in fibrillin genes (Robinson and Godfrey, 2000). Type-VI collagen microfibrils are found in most vertebrate tissues where they form an extensive microfibrillar network linking cells and many ECM components and maintaining the integrity of tissues such as blood vessels, lung, and skin (Kielty and Grant, 2002).
There is a pressing need to understand the effect of substrate on the conformation of native supramolecular assemblies to aid in the interpretation of solid-phase binding assay data and to gain insights into possible local charge environment dependent conformational switches in vivo. In this study, AFM in ambient conditions has been used to determine the influence of substrate on the conformation of isolated fibrillin and type-VI collagen microfibrils. We report marked substrate-dependent changes that have implications for functional analyses and provide new insights into microfibril supramolecular conformation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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Sample and substrate preparation
Adult bovine ciliary zonule and foetal bovine nuchal ligament microfibrils were isolated in native nondenaturing conditions by bacterial collagenase digestion and size exclusion chromatography using modifications of a previously described methodology (Kielty et al., 1991; Wess et al., 1998). Briefly, tissue was incubated with 0.5 mg/ml collagenase, hyaluronidase (final concentration, 5 units/ml), freshly prepared protease inhibitors (2 mM PMSF, 5 mM NEM), and 10 mM CaCl2 in column buffer (400 mM NaCl, 50 mM Tris-HCl, pH 7.4) for 18 h at 4°C. Samples were subsequently centrifuged at 5,000 g for 5 min, and the supernatant was size fractionated on a Sepharose CL-2B column in column buffer. The excluded volume (Vo) of ciliary zonule digests contained abundant fibrillin microfibrils. The excluded volume of nuchal ligament digests contained abundant fibrillin and type-VI collagen microfibrils.
Crosses were scored onto metal sample stubs using a single-edged razor blade. Trimmed mica sheets were adhered to the sample stubs using clear nail varnish. Cleaved mica surfaces were prepared immediately before use by peeling off the top layer of mica with adhesive tape. PLL-coated mica (mica-PLL) was prepared by 2-min incubations with 80 µl 0.01% w/v PLL, followed by three consecutive washes with 300 µl distilled H2O. Excess liquid was removed from the surface by capillary action at the edge, and the sample was allowed to air dry in a dust-free environment for 30 min before microfibril deposition. Glass coverslips were washed with ethanol followed by distilled H2O and allowed to air dry for 30 min before sample adsorption. Aliquots of microfibril-containing Vo were diluted 1:8 with column buffer. A sufficient volume of the diluted sample was pipetted onto the substrate and incubated for 1 min before three consecutive washes with 300 µl distilled H2O. Excess liquid was removed by capillary action and the prepared samples were allowed to air dry overnight in a dust-free environment before microscopical analysis.
The mica to mica-PLL substrate gradient was prepared using a freshly cleaved mica surface adhered to a sample stub marked with a central cross and radii marked at 1000-µm intervals. A 10-µl drop of PLL was pipetted onto the center of the cross, incubated for 2 min, washed three times with 300 µl distilled H2O, and air dried for 30 min. A sufficient volume of 10% v/v colloidal gold suspension was incubated on the substrate for 10 min before washing as previously described and air drying for 30 min. A sufficient volume of 1:8 dilution ciliary zonule Vo was incubated on the substrate for 1 min before washing and air drying overnight in a dust-free atmosphere.
AFM and image analysis
Samples were imaged by intermittent contact mode in air using a Multimode AFM with a Nanoscope IIIa controller and an E scanner (Veeco Instruments). The Nanoscope optical viewing system was used to aid laser alignment on the cantilever and to locate the cantilever in relation to the cross scored on the metal sample stub. The optical viewing system was calibrated using an electron microscope grid with 45 µm x 45 µm holes. Cantilever oscillation frequencies and drive amplitudes were determined by the Nanoscope software. Height images were captured at a scan rate of 1.2 Hz and scan sizes of 5, 3, 2.5, 2, or 1 µm. The set point was adjusted to just below the point at which tip-sample interaction was lost. The instrument was calibrated periodically using a grating with 180-nm deep, 10 µm x 10 µm depressions and during scans with 5 or 10 nm colloidal gold particles. All images were captured at a relative humidity of 30% ± 5%.
All measurements were conducted on height data, which was first-order flattened using the Nanoscope software. Curved filaments were straightened using the public domain program ImageJ (National Institutes of Health; available on the internet at http://rsb.info.nih.gov/ij/) and the straighten plug-in that fits a nonuniform cubic spline to user-supplied points (Kocsis et al., 1991). Extracted regions were scaled and axial height profiles determined using routines written in Microsoft Visual Basic 6.0 (Redmond, WA). Periodicity determinations were calculated from WSxM developer version 8.0 (Nanotec Electronica, Madrid, Spain; available on the internet at http://www.nanotec.es/). Power spectral analyses of type-VI collagen microfibril periodicities were performed on axial height profiles using WSxM. Roughness analyses (root mean square (RMS)) of PLL to mica binding were carried out with Microsoft Visual Basic 6.0 routines that used the equation employed by the Nanoscope software (Eq. 1) where Zave is the average Z-value within a given area, Zi is the current Z-value and N is the number of points within the given area.
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Substrate characterization
Mica, mica-PLL, and glass surface topologies were characterized by AFM. On some substrates RMS roughness values have been shown to be dependent on the measurement area (Long and Chen, 2001). RMS roughness values were calculated for a range of areas (linear dimensions from 31.25 nm x 31.25 nm to 1000 nm x 1000 nm) for 2 µm x 2 µm scans on each substrate, in triplicate.
The binding kinetics of PLL to freshly cleaved mica were investigated over a 32-min time course at time points of 0, 1, 2, 4, 8, 16, and 32 min. RMS values, of mica, mica-PLL, and glass, were calculated for 60 (105 x 105 nm) areas randomly placed over three 2 µm x 2 µm scans. The percentage coverage of PLL on the mica surface was calculated by autothresholding in ImageJ.
Mica, mica-PLL, and glass substrate hydrophobicity/hydrophilicity was quantified by measuring the drop contact angle (
), formed between the liquid-vapor and solid-liquid interfaces (Adamson, 1990; Bachmann et al., 2000). Drops of column buffer (10 µl) were pipetted onto the substrate; for drop sizes of 10 µl or less, gravity effects were not considered significant (Bachmann et al., 2000). Top and side view images were captured within 1 min using a desktop video camera. Sessile drop contact angles (appS) were determined from side-view images using Eq. 2 (Colwell et al., 2003) where H is the drop height and D the length of the section in contact with the surface.
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Advancing (appA) and receding (appR) contact angle measurements were performed using a modification of the sessile drop technique (Colwell et al., 2003). Incremental drops of column buffer (5-µl and 10-µl volumes for mica-PLL and glass, respectively) were pipetted onto the substrate. Images horizontal to the surface were captured at each increment. Receding contact angles were measured by the removal of 5- or 10-µl volumes of column buffer. All contact angle measurements were carried out at ambient laboratory relative humidity (30% ± 5%), in triplicate.
Drop spreading experiments were conducted on mica, mica-PLL, and glass substrates in triplicate. Drops of column buffer (5 µl) were pipetted onto the substrate, and the contact area of the spreading drop determined by the capture of 640 x 480 pixel movies at a frame rate of 3.75 frames s–1 over a 2-min time period. Drop radii were calculated from drop areas measured using ImageJ.
The influence of substrate on microfibril morphology
Adult bovine ciliary zonule or foetal bovine nuchal ligament microfibrils were adsorbed onto mica, mica-PLL, or glass substrates. Air-dried microfibrils were examined by AFM at multiple positions on each substrate. Consecutive bead/interbead repeats lying on clean substrates were extracted and straightened from 1 µm x 1 µm height scans. Individual repeats within the periodicity range 54.7–58.6 nm (29 pixels ± 1 pixel at a pixel size of 1.95 nm) were extracted and averaged.
To test the hypothesis that observed variations in microfibril morphology were substrate induced and not due to adhesion of morphologically distinct subpopulations, microfibril morphology was determined across a mica to mica-PLL interface by AFM. Gross displacements of the tip over the substrate were calculated from images captured from the Nanoscope optical viewing system calibrated against images of electron microscope grids (Athene old 400, 45 µm x 45 µm grid squares). AFM height images (3 µm x 3 µm) were captured at 12.15-µm intervals over a radial distance of 1100 µm starting at a radial position 50 µm < r0 (drop radius at 0 s) and ending at a radial position 50 µm > r120 (drop radius at 120 s). Gold particles were counted after thresholding in ImageJ. For height images with a relatively low number of adsorbed gold particles, mean RMS roughness values were calculated for three randomly placed areas (500 nm x 500 nm) within each scan. The interface between the mica and mica-PLL coated substrate was detected by an increase in surface Z-range (RMS) and the number of adhered negatively charged colloidal gold particles. Once the interface was located four (1 µm x 1 µm) AFM height scans were captured at 80-µm intervals spanning the interface region.
Hydropathy analysis
Mean hydropathy scores were calculated for amino acid sequences using the amino acid hydropathy values of Kyte and Doolittle (1982). Hydropathy profiles were calculated with a window size of 31 residues for the coding sequence following posttranslational modifications.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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appS = 21.2°, SD = 2.5°), mica-PLL less hydrophilic (appS = 43.2°, SD = 6.0°), and glass hydrophobic (appS = 90.3°, SD = 3.1°) at a relative humidity of 30% (Fig. 1, a–c (i)). Marked contact angle hysteresis was observed on mica-PLL substrates (Fig. 1 f). Contact angle hysteresis was less evident on the glass substrate (Fig. 1 g) and the hysteresis loop was closed.
The angle formed between the liquid-solid and liquid-vapor interfaces is called the wetting or contact angle (). Surfaces are defined as hydrophilic, or wettable, if < 90°; hydrophobic, or nonwettable, if 
90°, and perfectly wettable if is zero or nonmeasureable (Dury et al., 1998; Sweeney et al., 1993). On perfectly wettable surfaces, drops of water spread completely (Sweeney et al. 1993), on partially wettable (hydrophilic) or nonwettable (hydrophobic) surfaces, drops of liquid assume a shape, which depends on the relation between the free energies of the three involved surfaces. Experimentally, on nonideal surfaces, the measured contact angle (apparent contact angle; app) will be an average of microscopic contact angles (m) that follow local surface deformations (Dury et al., 1998). Although measurements are commonly made on a sessile drop (Bachmann et al., 2000), on most solid surfaces the measured contact angle as the drop advances (appA) across the surface is different from contact angle measured as the drop recedes (appR). This difference in advancing and receding contact angles, denoted as contact angle hysteresis (Schwartz and Garoff, 1985), may be induced by surface roughness or chemical heterogeneities (de Gennes, 1985).
Cleaving mica forms an atomically smooth chemically homogeneous substrate that would not be expected to exhibit contact angle hysteresis. The apparent contact angle of 21.2° indicates that the mica substrate was relatively wettable/hydrophilic. Glass substrates had advancing contact angles (appA) that decreased from 85° to 74° with increasing drop volume (Fig. 1 g), an effect that may be due to gravity (Bachmann et al., 2000). The relatively small contact angle hysteresis may be due to surface roughness, although some studies suggest that the effects of surface roughness on contact angle hysteresis are minimal when roughness is <100 nm (Neumann and Good, 1972; Extrand and Kumagai, 1997) and that surface chemistry is more important (Extrand and Kumagai, 1997). Glass and mica-PLL substrates are of comparable roughness but mica-PLL substrates exhibit large contact angle hysteresis effects (Fig. 1, f and g). We would suggest therefore, that roughness at the sub-nm scale does not explain the differences observed in hysteresis in these two systems. In general the contact angle of clean glass surfaces and water is close to 0° with glass hydrophobicity a consequence of the adsorption of organic substances from the atmosphere (Takeda et al., 1999). The mica-PLL system is chemically heterogeneous, and it is likely that the glass substrate is also chemically heterogeneous; therefore the differences in hysteretic behavior may be attributable to the extent and chemical nature of the heterogeneity. Kinetic hysteresis may dominate on mica-PLL substrates as the adsorbed PLL molecules swell or reorganize in the liquid phase (Morra et al., 1990). The spreading behavior and apparent sessile drop contact angles suggest that each of the three substrates is partially wettable in the system substrate-sodium chloride solution air, although the degree of wettability decreases in the order mica > mica-PLL > glass.
Mica minerals are layered crystals in which tetrahedral sheets of (Si, Al)2O5 are ionically linked by a central Al2(OH)2 layer (Bailey, 1984). In the uncleaved muscovite mica crystal the net negative charge of basal oxygens is balanced by a layer of hexagonally coordinated K+ cations. Following cleavage, this layer is completely neutralized by the negative aluminosilicate lattice (Gaines and Tabor, 1956). In water some of the K+ ions dissociate from the surface resulting in a negative surface charge density of 15,000 negative charges/µm2 at neutral pH (Pashley, 1981; Muller et al., 1997) presenting a hydrophilic surface (Kowalewski and Holtzman, 1999). Subsequent microfibril adsorption experiments were carried out in a salt solution. Microfibril preparations were eluted from the size exclusion column in a 400 mM NaCl solution buffered to a pH of 7.4 by 50 mM Tris-HCl and 10 mM CaCl2 was present in the collagenase digestion buffer. In salt solutions, the extent of the dissociated and replaced K+ ions from the mica surface depends on the electrolyte concentration (Gaines and Tabor, 1956; Pashley, 1981). The surface charge in solution is balanced by bound counterions that form the Helmholtz layer and a diffuse charge cloud, the electric double layer (Muller et al., 1997). Cations bind to the negatively charged surface rendering the mica positively charged (Forbes et al., 2001). Therefore, during adsorption of microfibrils to mica the surface was hydrophilic and positively charged. Previous studies have demonstrated that adsorbed PLL reverses the negative charge of mica and silica (Luckham and Klein, 1984; Velegol and Tilton, 2001). The adsorption of negatively charged colloidal gold in the absence of divalent cations also suggests that PLL reverses the charge of mica. The influence of Helmholtz and electric double layers would be reduced on a positively charged surface. The low wettability of glass coverslips may be due to adsorption of atmospheric substances by surface OH groups (Takeda et al., 1999). From our data we conclude that microfibrils adsorbing to glass will encounter a hydrophobic microenvironment and mica-PLL will present a less hydrophilic microenvironment than mica alone.
Substrate-dependent morphology of fibrillin microfibrils
Fibrillin microfibrils isolated from foetal bovine nuchal ligament were identified by their unique repeating beaded structure and untensioned periodicity of 
56 nm on mica, mica-PLL, and glass substrates, (Fig. 2, a–c) (Sherratt et al., 2001). Fibrillin microfibrils adsorbed on the hydrophilic mica substrate had a diffuse appearance (Fig. 2 a). Microfibrils adsorbed to the less hydrophilic mica-PLL substrate appeared more condensed with a shoulder region (bead asymmetry), which conferred an orientation to the microfibril (Fig. 2 b). Morphologically indistinguishable microfibrils were observed on glass substrates (Fig. 2 c). Similar variations in microfibril morphology have been reported for microfibrils visualized by rotary shadowing transmission electron microscopy (TEM) (Wright and Mayne, 1988; Kielty et al., 1991). Although the removal of aqueous solvent during air drying may affect microfibril morphology, recent studies have demonstrated that the major structural features identified by electron microscopy and AFM in dehydrated microfibrils such as the bead/interbead structure, periodicity, and shoulder region are also present in fully hydrated microfibrils in solution (Sherratt et al., 2003). We suggest that the charge on mica has a profound effect on microfibril morphology and that mica-PLL and glass substrates induce fewer morphological rearrangements on adsorption.
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). Given the reported variability of microfibril composition, the diffuse and condensed morphologies observed on mica and mica-PLL/glass substrates, respectively, could result from the substrate-specific adsorption of compositionally and hence morphologically distinct subpopulations. To test this hypothesis, microfibrils were adsorbed onto a mica substrate with a central PLL-coated region. The PLL drop spread over the mica surface during the 2-min incubation period (Fig. 3, a and b). Drop spreading provides a mechanism for the creation of a PLL gradient. At the radial position r0 the mica substrate as exposed to PLL for 2 min, between radial positions of 3500–3807 µm there was an approximately linear decrease in mica to PLL contact time (Fig. 3 b). Gold particles required a critical amount of adsorbed PLL to adhere to the substrate (Fig. 3 d), but the amount of adsorbed PLL decreased in a linear fashion after this point as determined by RMS roughness measurements (Fig. 3 e). The position of the mica-PLL/mica interface was located by the adsorption/nonadsorption of 10-nm diameter negatively charged colloidal gold spheres that only bound to the positively charged PLL (Fig. 4 b). If the observed morphologies were substrate induced then microfibrils with an intermediate morphology would be observed at the interface. At positions flanking the interface, fibrillin microfibril morphologies were similar to those observed previously, diffuse on mica (Fig. 4 d (iv)) and condensed with a pronounced shoulder region on mica-PLL (Fig. 4 d (i)). Fibrillin microfibrils at positions ii and iii, which lie within the mica to mica-PLL interface (Fig. 4 d (ii and iii)), had an intermediate morphology with a shoulder region that was more pronounced than on mica (Fig. 4 e (i)) but less pronounced than on mica-PLL (Fig. 4 e (iv)). The calculated axial slope within this plateau region increased between positions i–iv (Fig. 4 f). We conclude that distinct microfibrillar morphologies are substrate induced rather than substrate selected.
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). Hydrophilic substrates may disrupt interdomain ionic interactions inducing an extended fibronectin morphology and a diffuse fibrillin microfibril morphology. Sample-substrate interactions on hydrophobic substrates are thought to be dominated by relatively weak van der Waals forces and hydrogen bonds (Betley et al., 2001). These weak interactions may not interfere with intermolecular ionic interactions reducing substrate-induced distortion of the sample. As with fibronectin (Bergkvist et al., 2003) and synthetic dendrimers (Betley et al., 2001) the morphology of fibrillin microfibrils on mica-PLL and glass may more closely resemble the morphology in solution than fibrillin microfibrils adsorbed to mica.
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; Hanssen et al., 1998; Baldock et al., 2001). The disruption of the shoulder region on mica substrates identifies an axially located region of the microfibril, which is particularly sensitive to local charge environments. Although the amino acid sequence of fibrillin-1 is predominantly hydrophilic (Fig. 5 b), there are no extended regions of increased hydrophilicity. Fibrillin-1 is a glycoprotein with 12 potential N-glycosylation sites. N-linked glycosylation of recombinant putative glycosylation sites within fibrillin-1 peptides has been demonstrated by shifts in electrophoretic mobility following endoglycosidase H treatment (Ashworth et al., 1999). Complex N-linked oligosaccharides are negatively charged if they contain sialic acid. Many of the glycosylation sites are within a section of the monomer, which is predicated to lie within the interbead region within the microfibril (Baldock et al., 2001). Interactions between hydrophilic N-linked oligosaccharides and the hydrophilic mica may cause structural rearrangements that disrupt the shoulder region. Additional regions of increased hydrophilicity may result form the heteropolymeric nature of fibrillin microfibrils. TEM and scanning TEM ultrastructural approaches including cuprolinic blue staining (Kielty et al., 1996) and mass changes following enzymatic digestion (Sherratt et al., 1997) provided strong evidence for an association of chondroitin sulfate proteoglycans (CSPGs) with the fibrillin microfibril bead. A small sulphated CSPG interacts with fibrillin-1 at a site within or adjacent to the proline-rich region (Trask et al., 2000). CSPGs consist of a core protein and one or more highly negatively charged glycosaminoglycan (GAG) side chains. The structure of a shoulder region composed of associated CSPG and hydrophilic GAGs may be disrupted by a hydrophilic substrate. Substrate-dependent interactions have been demonstrated between fibrillin-1 fragments and microfibril associated molecules, tropoelastin, and MAGP-1 (Rock et al., 2004).
Isolated microfibrils have been shown to support the attachment of vascular smooth muscle cells (Kielty et al., 1992) and cell-specific integrins 
5ß1 and vß3 induce cell spreading, alter cytoskeletal organization, and upregulate fibrillin-1 synthesis (Bax et al., 2003). Substrate-induced conformational changes may alter cellular recognition of fibrillin microfibrils by RGD (Arg-Gly-Asp) site masking or by relative movement of the RGD and potential synergy sites (Krammer et al., 2002). The RGD site is located within the interbead region in close proximity to a number of N-linked glycosylation sites (Fig. 5 b). Microfibril-associated molecules such as MAGP-1, tropoelastin, and fibulin-2 are thought to bind at or near the proline-rich region (Rock et al., 2004). Substrate-induced changes in GAG conformation may mediate the binding of associated molecules. Local charge environments may also affect fibrillin microfibril morphology in vivo. The vertebrate elastic fiber is composed of fibrillin microfibrils and highly hydrophobic elastin (Swiss-Prot P15502; hydropathy score = 0.706). It is not clear whether elastin-associated microfibrils differ morphologically and functionally from those that do not associate with elastin (Kielty et al., 2002a). We suggest that local charge environment conformational changes may also occur in vivo acting as a molecular switch to hide or reveal binding sites in compositionally identical elastin-associated and nonelastin-associated fibrillin microfibrils.
Substrate-dependent morphology of type-VI collagen microfibrils
Type-VI collagen microfibrils have a unique and complex repeating double-bead structure with an untensioned periodicity of 104–112 nm (Ball et al., 1999; Baldock et al., 2003). Beads are composed primarily of the noncollagenous N- and C-terminal regions and the interbead of four collagen triple helical regions (Furthmayr et al., 1983; Baldock et al., 2003). Type-VI collagen microfibrils isolated from foetal bovine nuchal ligament had well-defined double-bead and interbead regions when adsorbed to a mica-PLL substrate (Fig. 2 b). As with fibrillin microfibrils, morphologically indistinguishable assemblies were observed on mica-PLL and glass substrates (Fig. 2 c). Beads had a maximum height of 3 nm and interbeads were 1 nm in height (Fig. 6 a). Type-VI collagen microfibrils, isolated from foetal bovine nuchal ligament and aorta, with similar bead/interbead heights and repeating periodicities have previously been identified on PLL-coated mica substrates by AFM (Baldock et al., 2003). Adsorption to a mica substrate induced major morphological changes (Figs. 2 a and 6 b). The characteristic double-bead/interbead structure was largely absent and periodicity could not be determined visually (Fig. 6 b). These structures were identified as type-VI collagen microfibrils using a number of observations. In regions of the mica surface where these structures formed bundles, double beads were clearly identifiable (Fig. 2 a (ii)). Double beads with a similar height maximum to mica-PLL double beads were occasionally observed on regions of the microfibril exposed to bare mica (Fig. 6 b). Power spectral density analyses of these structures revealed an underlying periodicity very similar to the reported periodicity of type-VI collagen microfibrils and the periodicity measured on mica-PLL (Figs. 6, c and d). Unlike the region-specific conformational changes observed for fibrillin microfibrils, type-VI collagen microfibrils respond to more hydrophilic environments with major structural rearrangements along the whole repeating unit.
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3(VI) N9–N1 = –0.01). The triple helical region is highly hydrophilic and the RGD sites are clustered in this region. Color-coding domains according to mean hydropathy revealed that most of the noncollagenous domains are slightly hydrophilic (3(VI) N1–N4 mean = –0.198) but 3(VI) domains (N5–N9) are slightly hydrophobic (mean = 0.139). The current folding model of the type-VI collagen N-terminal globular region predicts that these domains will form a loop located on one side of the N-terminal globular region (Specks et al., 1992; Baldock et al., 2003). In the assembled microfibril, these hydrophobic faces may come into contact forming a hydrophobic pocket, which helps to stabilize the microfibril. It is unlikely that these hydrophobic interactions play a role in microfibril assembly as 3(VI) N5–N9 domains are not needed for microfibril assembly (Lamande et al., 2002). We propose that the difference in morphology is due to the higher exposure of charged groups on the microfibril to the charged hydrophilic mica. In contrast to the localized charge within fibrillin microfibrils, the highly charged triple helical region of type-VI collagen spans the whole repeat (Baldock et al., 2003). The majority of the globular domains in type-VI collagen have homology to von Willebrand factor (vWF) A domains and vWF itself has been shown to be sensitive to the local surface charge environment. On a modified hydrophobic glass surface, adsorbed vWF appeared compact by AFM, on a hydrophilic mica surface, vWF extended to cover a greater surface area (Raghavachari et al., 2000).
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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The increasing reliance on solid-phase binding assays to investigate protein-protein and protein-cell interactions highlights the importance of physisorption effects on protein function. Local charge environments may also induce conformational changes in supramolecular assemblies in vivo. Elastin-associated fibrillin microfibrils may differ from nonelastin-associated microfibrils only in their charge environment induced conformation and not in composition.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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M.J.S. and C.M.K. acknowledge the support of the Medical Research Council (UK). D.F.H. acknowledges the support of the Wellcome Trust.
Submitted on September 3, 2003; accepted for publication January 29, 2004.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSACKNOWLEDGEMENTSREFERENCES |
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