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School of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT UK
Reprint requests to: Prof. Sheena E. Radford, School of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT UK; e-mail: s.e.radford{at}leeds.ac.uk; fax: 44-113-233-3167.
(RECEIVED January 31, 2001; FINAL REVISION May 17, 2001; ACCEPTED May 29, 2001)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1101/ps.4901.
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Abstract |
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10 nm wide, but are shorter and assemble more rapidly than those produced from the oxidized protein. These data show that population of non-native conformers of ß2m at neutral pH by reduction of its single disulphide bond is not sufficient for amyloid formation. Instead, association of one or more specific partially unfolded molecules formed at acid pH are necessary for the formation of ß2m amyloid in vitro. Further experiments will now be needed to determine the role of different oligomeric species of ß2m in the toxicity of the protein in vivo. Keywords: ß2-Microgobulin; amyloidosis; disulphide bond; protein fibril
Abbreviations: ß2m, human ß2-microglobulin • thio-T, thioflavin-T • ANS, 1-anilinonapthalene-8-sulphonic acid • DRA, dialysis-related amyloidosis • ESI MS, electrospray ionisation mass spectroscopy • GuHCl, guanidinium chloride • red-ß2m, ß2m in which the single disulphide bond is reduced • ox-ß2m, ß2m in which the single disulphide bond is oxidized
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Introduction |
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20 proteins are known to be associated with human amyloid (Sunde et al. 1997). Even though the structure of the soluble form of the known human amyloidogenic proteins varies widely and includes 
-helical, ß-sheet, and mixed /ß proteins, the fibrils they produce exhibit similar structural characteristics. Electron microscopy (Sunde and Blake 1997; Jimenez et al. 1999), dye-binding assays (Naiki et al. 1989; Klunk et al. 1999), and X-ray diffraction studies (Sunde et al. 1997) have established that amyloid fibrils are long, unbranching protein fibers with a cross-ß structure.
Dialysis-related amyloidosis (DRA) involves the aggregation of ß2-microglobulin (ß2m) into amyloid deposits and is a serious complication of long-term haemodialysis (Gejyo et al. 1986). ß2m is a small (99-residue) protein that has a seven-stranded ß-sandwich fold typical of proteins belonging to the immunoglobulin superfamily (Bjorkman et al. 1987) (Fig. 1
). The two ß-sheets are held together by a single buried disulphide bond that links Cys25 and Cys80 in ß-strands 2 and 6, respectively. The disulphide bond is highly conserved in the immunoglobulin superfamily and stabilizes the native fold of these proteins (Isenman et al., 1975). In vivo, ß2m is continuously shed from the surface of cells displaying MHC class I molecules. It is then carried in the plasma to the kidneys where it is degraded and excreted. As a consequence of renal failure, the concentration of ß2m in the plasma increases 25– to 35-fold (Gejyo et al. 1986), which ultimately leads to the deposition of the protein into amyloid fibrils in the musculoskeletal system (Ritz and Zeier 1996). Full-length wild-type ß2m, as well as modified and truncated versions of the protein, have been found in ß2m fibrils ex vivo, although no natural mutations of the gene sequence have been associated with the disease (Floege and Ehlerding 1996; Esposito et al. 2000). The role of sequence modifications in the mechanism of amyloidosis of ß2m in vitro and the influence of these and other factors in the development of the disease in vivo are currently poorly understood.
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10 nm wide, short (50–200 nm), and have a curvilinear morphology (McParland et al. 2000). On further acidification (to pH 1.6), the fibrils formed have the same width and morphology as those produced at pH 3.6, but extend to greater than 600 nm in length. Under these conditions a second acid-denatured species is generated that is less structured than partially unfolded ß2m at pH 3.6 (McParland et al. 2000).
The data obtained so far are consistent with a model for ß2m fibrillogenesis in vitro in which one or more partially unfolded molecules of the protein associate to form ordered fibrillar assemblies (McParland et al. 2000). To test this hypothesis further and to examine the role of protein stability in ß2m amyloidosis, we performed a study of the amyloidogenic properties of reduced ß2m (red-ß2m) in vitro, in which the single disulphide bond linking Cys25 and Cys80 has been disrupted (Fig. 1
). We show that red-ß2m is non-native even at neutral pH. Nevertheless the protein does not form amyloid fibrils under these conditions. Like its oxidized counterpart, red-ß2m forms fibrils at acidic pH and high ionic strength. The data indicate that destabilization of ß2m by reducing its disulphide bond is not sufficient for amyloidosis, but association of specific partially unfolded molecules formed at acidic pH are necessary for amyloidosis of this protein in vitro.
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Results |
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Characterization of the conformational properties of red-ß2m as a function of pH was performed using a variety of spectroscopic techniques. The far UV CD spectra of ox-ß2m and red-ß2m at pH 7.0 are shown in Figure 2A. Ox-ß2m at pH 7.0 and 10°C is native (Okon et al. 1992; McParland et al. 2000). The far UV CD spectrum of this species is characteristically weak in intensity, as is often observed for ß-sheet proteins (Woody 1995). Positive bands at 270 and 290 nm are observed in the near UV CD spectrum of ox-ß2m at pH 7.0, consistent with the packing of aromatic rings in a fixed conformation in the native protein (Fig. 2D). On reduction of the disulphide bond, there is a large increase in intensity of the negative ellipticity in the far UV CD, suggesting that ß-sheet secondary structure persists in red-ß2m at neutral pH (Fig. 2A). In addition, there is a small, but significant, decrease in intensity of the near UV CD spectrum on reduction of the protein at pH 7.0, suggesting that the tertiary structure of the reduced protein is altered relative to that of its native, oxidized counterpart (Fig. 2D). Analytical ultracentrifugation studies showed that although the oxidized protein is monomeric at pH 7, the reduced protein forms a mixture of monomers and tetramers at this pH (Table 1). Changes in tertiary structure (involving aromatic rings buried in the core of the protein), or quarternary structure (for example, involving Trp 60, which is solvent exposed in native oxidized ß2m), could account, at least in part, for the differences in the far UV CD spectra of the oxidized and reduced proteins at neutral pH. Nevertheless, the data show that red-ß2m is non-native at neutral pH.
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max. In contrast, ox-ß2m does not bind ANS under these conditions (Fig. 3A). These data suggest that red-ß2m at pH 7 retains significant secondary and tertiary structure but exposes non-native hydrophobic surface area, consistent with the population of non-native species in the reduced protein under these conditions.
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). Ox- and red-ß2m bind ANS at pH 3.6, indicating that a significant proportion of both species expose hydrophobic surface area at this pH (Fig. 3B). Ultracentrifugation studies indicate that both ox- and red-ß2m at pH 3.6 are polydispersed in the mixed buffer used (see Materials and Methods), forming a number of species ranging from monomers to large oligomers (Table 1). Because the oxidized and reduced proteins are polydisperse at this pH under the conditions used (Table 1), a more detailed interpretation of the CD spectra in terms of the nature of residual secondary structure present in each protein is not possible.
When both ox- and red-ß2m are acidified to pH 1.5, further denaturation of the proteins occurs, as judged by far and near UV CD (Fig. 2C,F). The far and near UV CD spectra of red-ß2m at pH 1.5 suggest that the protein retains residual secondary structure, but lacks significant fixed tertiary interactions under these conditions. At this pH, the oxidized protein has a similar near UV CD spectrum to its reduced counterpart above 260 nm, but has a distinct far UV CD spectrum, suggesting that the conformational properties of the two proteins differ in detail at this pH. Both ox-ß2m and red-ß2m bind ANS at pH 1.5 (Fig. 3C), indicating that both proteins expose hydrophobic surface area at this pH. Ultracentrifugation studies demonstrated that both ox-ß2m and red-ß2m form specific trimers at pH 1.5 (Table 1).
The amyloidogenic properties of red-ß2m
The above data indicate that reducing the disulphide bond of ß2m destabilizes the protein such that non-native species are populated at neutral pH. To investigate the role of these, and other partially unfolded species in amyloidosis in vitro, ox-ß2m and red-ß2m were incubated at different pH values and fibrillogenesis was initiated by the addition of 0.4 M NaCl. The structural properties of the protein were measured immediately after the addition of NaCl by far UV CD (Fig. 4), and the presence of fibrils was determined after incubation under these conditions for three days using thio-T binding and negative stain EM (Figs. 5 and 6). Increasing the ionic strength has little effect on the far UV CD spectra of ox-ß2m at pH 7.0 and pH 3.6, and red-ß2m at pH 7.0 (Fig. 4A,B). In contrast, the addition of NaCl to red-ß2m at pH 3.6 causes a rapid decrease in intensity in the far UV CD (Fig. 4B), which occurs concomitantly with the production of material that scatters light (measured by optical density; data not shown). At longer incubation times under these conditions, insoluble material that sedimented in the CD cuvette was produced. Both ox-ß2m and red-ß2m appear to partially refold on the addition of NaCl at pH 1.5 (Fig. 4C). Therefore, the addition of NaCl to ox-ß2m at pH 1.5 results in partial refolding of the molecule such that a species with a CD spectrum similar to that of acid unfolded ß2m at pH 3.6 is formed (Fig. 4B,C) (McParland et al. 2000). In addition, the decrease in intensity of the far UV CD signal of red-ß2m at <215 nm on the addition of NaCl is consistent with reformation of secondary structure in this partially unfolded state.
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; Table 1) (McParland et al. 2000). Under these conditions, the fibrils formed are 10-nm wide and have a curvilinear morphology (Fig. 6C,D). The fibrils formed from the oxidiszed protein at pH 3.6 are relatively short (ranging from 50–200 nm in length), whereas those formed at pH 1.5 have the same width and morphology as those produced at higher pH, but extend to >600 nm in length (Table 1) (McParland et al. 2000). Interestingly, incubation of red-ß2m in 0.4 M NaCl results in the formation of species capable of binding thio-T over the entire pH range studied (pH 8.0–1.0) (Fig. 5). Despite examining a large number of grids, fibrillar material was only observed by negative stain EM in samples incubated below pH 4.0 (Fig. 6A,B). At these pH values, curved fibrils with a diameter of 10 nm are formed. Fibrils of the reduced protein are shorter in length than those formed from ox-ß2m under the same growth conditions (Table 1). At pH 4.5, the formation of amorphous aggregates from red-ß2m results in light scattering that accounts for the apparent large increase in thio-T fluorescence at this pH. Because red-ß2m at neutral pH does not form high molecular-weight oligomers (as judged by ultracentrifugation, light scattering, and negative stain EM), binding of thio-T to samples of red-ß2m incubated above pH 4.5 most probably reflects an interaction of the dye with the tetrameric species formed from the reduced protein at this pH. At pH 7, ox-ß2m does not form tetrameric species and no thio-T binding is evident.
Fibril morphology is dependent on the rate of fibril growth
To examine the influence of growth rate on fibril length, the kinetics of fibrillogenesis of ox–ß2m and red-ß2m at pH 7.0, pH 3.6, and pH 1.5 were measured using a continuous thio-T binding assay (see Materials and Methods). At pH 7.0, monomeric ox-ß2m does not bind thio-T, consistent with previous observations that the native protein is not amyloidogenic (Fig. 5; Table 1) (McParland et al. 2000). Conversely, red-ß2m shows a significant rate of increase in thio-T fluorescence at pH 7.0, which presumably reflects the formation of the small oligomeric species detected by ultracentrifugation (see above and Table 1). Most interestingly, there is a correlation between the rate of fibril growth and fibril length (Table 1). Conditions that favor rapid growth (red-ß2m in 0.4 M NaCl at pH 3.6) result in the formation of numerous short fibrils, whereas those resulting in slower growth (ox-ß2m at pH 1.5) produce fewer, longer fibrils. Fibril length is also independent of the redox state of ß2m (fibrils of comparable length are formed from ox-ß2m at pH 3.6 and red-ß2m at pH 1.5). The observation that the length of fibrils increases when the association rate is lowered has long been established for a number of assembling systems (Whitesides et al. 1991; Bowden et al. 2001). Despite the apparent independence of fibril morphology on the redox state of the precursor protein, however, there is no reason to assume that the mechanism of ß2m amyloid formation from the oxidized and reduced proteins will also be similar. Further experiments will be needed to determine the nature of the nucleus and the mechanism of elongation of the oxidized and reduced proteins.
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Discussion |
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TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
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cDNA is associated with a human light chain amyloid disease (Perfetti et al. 1998). In this paper we have shown that reducing the single disulphide bond of human ß2m results in the destabilization of the native protein, such that non-native species are populated at neutral pH. Despite this, only partially unfolded species formed below pH 4.0 are capable of forming amyloid in vitro under the conditions investigated here. These results indicate that destabilization of native ß2m by reduction of its disulphide bond is not sufficient for the formation of amyloid in vitro. Instead, one or more partially unfolded conformers formed at low pH appear to be necessary for amyloidosis of ß2m in vitro. The symptoms of DRA are known to be brought about by the stimulation of macrophages to produce bone-resorbing cytokines, such as interleukin 1ß, tumor necrosis factor-, and interleukin 6, resulting in bone degradation (Hou et al. 2001b). It is also known that ß2m modified with advanced glycation end products may induce a local inflammatory response associated with DRA (Hou et al. 2001a). Whether the immature or mature fibrils of ß2m are responsible for different facets in the evolution of DRA is currently unknown. Moreover, it is also possible that soluble oligomeric species of ß2m, such as the trimeric and tetrameric species observed here under different conditions, could also be involved. In accord with this view, studies of other proteins have indicated that nonfibrillar species could be the culprits of these types of disease (for review, see Lansbury 1999).
The characteristics of the fibrils formed from ox-ß2m and red-ß2m depend on the pH at which they are formed. Although fibrils of both ox-ß2m and red-ß2m formed in vitro have a common highly curved morphology and are 10-nm wide, the length of fibrils produced depends on the initial rate of their formation but is independent of the redox state of the protein. Fibrils formed from both ox-ß2m and red-ß2m at acidic pH in vitro are shorter and more curved in morphology than those formed from most other proteins (Brancaccio et al. 1995; Goldsbury et al. 1997; Sunde and Blake 1997). ß2m fibrils have also been formed in vitro by extension of amyloid fibrils from patients with DRA with full-length ß2m (Naiki et al. 1997), as well as with ß2m truncated by six residues at its amino terminus (Esposito et al. 2000). Under such conditions, the fibrils formed are 10-nm wide, but are longer and straighter than those formed de novo in vitro. The latter may therefore represent immature fibrils or, in the case of the very short fibrils formed from red-ß2m at pH 3.6, early assembly intermediates. Interestingly, however, further assembly of the curved immature fibrils formed in vitro to the mature fibrils observed ex vivo has not been observed to date, despite exploring a range of conditions (Naiki et al. 1997; McParland et al. 2000). Consistent with these results, incubation of immature fibrils assembled from both ox-ß2m and red-ß2m de novo in vitro at pH 3.6 in 0.4 M NaCl for up to 8 mo at 37°C have not yielded mature fibrils akin to those formed in vivo. Further experiments will be needed to determine the factors that influence the extent of ß2m fibril assembly in vitro, the role of the curved immature fibrillar species in the assembly of mature amyloid fibrils and the influence of these different species in the evolution of the pathogenicity of ß2m amyloid in vivo.
Implications for ß2m amyloidosis in vivo
The incidence of dialysis related amyloidosis in patients receiving haemodialysis for more than two decades is 100% (Floege and Ehlerding 1996). Studies of the epidemiology of haemodialysis-related amyloidosis offer clues as to its possible causes. Fibrils formed from ß2m in vivo are often located in areas rich in collagen, such as the joints (Gejyo et al. 1995; Floege and Ehlerding 1996), resulting in the crowding of macrophages in these areas (Ohashi et al. 1992; Ayers et al. 1993). ß2m amyloid has been observed directly within macrophage lysosomes (Van Ypersele and Drucke 1996), raising the possibility that uptake of the protein into lysosomes may be involved in the development of ß2m amyloidosis. In contrast with the neutral pH of serum, the acidic pH within lysosomes would promote partial unfolding of ß2m to species capable of assembly into amyloid fibrils. In addition, it has been discovered recently that the enzyme gamma-interferon-inducible lysosomal thiol reductase is responsible for the enzymatic reduction of disulphide bonds in lysosomes (Arunachalam et al. 2000), raising the possibility that reduction of disulphide bonds could be involved the initiation of protein aggregation within this cellular compartment.
Native monomeric ß2m has been recovered from amyloid deposits of patients with DRA (Gejyo et al. 1986; Campistol et al. 1996; Bellotti et al. 1998). Analysis of this material has suggested that at least a significant proportion of the protein in amyloid deposits ex vivo is in the oxidized form. In addition, the observation that amyloid can be formed from immunoglobulin light chains (Helms and Wetzel 1996; Raffen et al. 1999), lysozyme (Booth et al. 1997), insulin (Bouchard et al. 2000), and ß2m (Connors et al. 1985; Ono and Uchino 1994; Naiki et al. 1997; Bellotti et al. 1998; McParland et al. 2000) in their oxidized forms suggests that reduction is not a prerequisite for amyloidosis. The rapid formation of small oligomers or immature protofibrils of red-ß2m within lysosomes could seed polymerization of partially unfolded ox-ß2m in the acidic pH environment therein. Assembly of truncated ß2m into fibrils from such seeds would also be possible in the neutral pH of serum (Esposito et al. 2000). Further experiments will be needed to elucidate the role of disulphide bond reduction in ß2m amyloidosis in vivo. Nevertheless, the results presented here indicate that population of partially unfolded conformers of ß2m in an acidic environment are important features of ß2m amyloidosis in common with many other human amyloid diseases (Lai et al. 1996; Swietnicki et al. 1997; Guijarro et al. 1998).
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Materials and methods |
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ß2m overexpression and purification
The gene encoding human ß2m was obtained from the plasmid BJ192 (kindly provided by B. Jakobson, Institute of Molecular Medicine, Oxford, UK). The gene was subcloned into the vector pET23a (Promega), and the resulting plasmid pINKwt was transformed into BL21(DE3) cells. The cells were grown in LB media containing 0.25 mg/ml carbenicillin and protein expression was induced by the addition of 1 mM IPTG when the optical density had reached an OD600 of 0.6 (McParland et al. 2000). After 12 h, the bacteria were harvested. Overexpressed ß2m was sequestered in the cells as inclusion bodies. These were isolated and the protein purified as described in McParland et al. (2000). A yield of 35 mg pure ß2m/L culture was obtained. The protein was shown to be >95% pure by SDS PAGE and of the expected molecular weight by ESI MS (expected Mr=11,860, observed Mr=11,860±0.72). The protein was stored at -20°C as a lyophilized powder.
Preparation and analysis of reduced ß2m
Lyophilized ß2m was dissolved at room temperature in 25 mM Tris HCl containing 6 M GuHCl, 10 mM DTT, pH 8.0. After 1 h at room temperature, the sample was cooled to 4°C and the protein was purified by rapid gel filtration using Pharmacia PD10 columns equilibrated with 10 mM Tris HCl, 10 mM DTT, pH 8.0. Any precipitate was removed by centrifugation and the sample was then passed through a 0.2 µM filter and used immediately. To determine the extinction coefficient of the reduced protein, the protocol was repeated using a known amount of ox-ß2m at a protein concentration of 0.2 mg/ml. Under these conditions, no precipitate was formed on removal of the denaturant. The extinction coefficient of reduced refolded ß2m at 280 nm in 10 mM Tris HCl, 10 mM DTT, pH 8.0 was determined to be 26,827 M-1cm-1. The extinction coefficient of the oxidized protein under these conditions was taken as 19,850 M-1cm-1 (Berggard and Bearn 1968).
To demonstrate that ß2m was fully reduced, two assays were performed. First, the sample (0.2 mg/ml in 25 mM Tris HCl, 25 mM sodium acetate, 25 mM glycine, 25 mM MES, 10 mM DTT, pH 8.0) was modified with iodoacetic acid. After an initial 15-min incubation at 25°C with 100 mM iodoacetate at pH 8.0 (to modify the DTT in the sample buffer), the protein was unfolded by the addition of 6 M GuHCl and the sample was incubated in the presence of iodoacetate for a further 20 min. The sample was then refolded by 10-fold dilution into 10 mM Tris HCl buffer, pH 8.0, dialyzed against water and the mass of the protein determined by ESI MS. The majority of the ox-ß2m was not modified by iodoacetic acid (observed Mr=11,860±1.5). A fraction of molecules (25%), however, contained modification of a single surface residue (Mr=11,918.5±0.53), possibly the solvent exposed histidine, H13. Red-ß2m was fully reduced using this protocol, 62% of molecules containing two carboxymethyl moieties (Mr=11,978±0.64). The remainder showed modification of a third residue (Mr=12,035±3.50). Repeating the experiment on the reduced, refolded protein after storage at a range of pH values (1.5–8.0) at 37°C for up to 6 days under an atmosphere of N2 showed that no re-oxidation of the two Cys residues had occurred.
In the second assay, ox-ß2m and red-ß2m were analyzed by reverse-phase HPLC using a C18, 300Å column (Brownlee Aquapore OD-300 100 mmx2.1 mm). All HPLC separations used an acetonitrile-H2O gradient in the presence of 0.1% trifluoroacetic acid (TFA). Comparisons of elution times were used to ascertain the redox state of the protein. Peaks were identified by their relative intensities. The gradient used for the final separations was A: 0.1% TFA, B: 0.1% TFA/90% CH3CN; 0 min 10% B, 2 min 10% B, 7 min 30% B, 37 min 60% B, 45 min 90% B, 47 min 90% B, 52 min 10% B. Under these conditions, the retention time of ox-ß2m and red-ß2m were 28.7 min and 40.8 min, respectively. The data confirmed that the protein was fully reduced using the procedure outlined above.
Ultracentrifugation
Sedimentation velocity experiments were carried out using a Beckman Optima XL-I analytical ultracentrifuge (Beckman, Palo Alto, CA) fitted with a scanning absorption optical system. To determine the sedimentation coefficients of red-ß2m, the protein was prepared as described above. Double-sector cells were filled with 280 µl of 0.2 mg/ml protein solution and 300 µl of reference solvent (25 mM Tris HCl, 25 mM sodium acetate, 25 mM glycine, 25 mM MES, 10 mM DTT) in their respective channels. Solutions were run at 60,000 rpm at 13°C and solute distributions were scanned at 280 nm. A series of 51 scans were taken at fixed time intervals over 2.5 h. Scans were analyzed using the program XLAVEL (Colfen et al. 1996), giving the sedimentation coefficient for each protein.
Thioflavin-T binding studies
Aliquots of red-ß2m in 10 mM Tris HCl buffer, pH 8.0 were diluted into buffers consisting of 25 mM Tris HCl, 25 mM sodium acetate, 25 mM glycine, 25 mM MES, and variable concentrations of NaCl to adjust the ionic strength at each pH to 400 mM. The samples were then passed through a 0.2 µM filter and diluted into the desired final solution conditions. A final ß2m concentration of 0.2 mg/ml was used throughout. Protein was incubated at 37°C for 72 h without stirring, after which time the samples were diluted 100-fold into the assay buffer (10 µM Thio-T, 50 mM Tris HCl, pH 8.5) before measurement. Thio-T fluorescence was measured using a PTI Quantamaster C-61 spectrofluorimeter set at 444 nm (excitation) and 480 nm (emission) using slit widths of 1 nm and 10 nm, respectively. The excitation and emission wavelengths were determined as peak values by performing excitation and emission scans in the presence of preformed ß2m fibrils. Control experiments showed that fibrils formed at lower pH values were stable at pH 8.5 over the time course of the measurement (10 sec). The florescence of thio-T in the absence of protein was also taken and used as a blank. Positive controls were included in all experiments using pre-formed ox-ß2m fibrils. The data were scaled by taking the thio-T fluorescence value of the sample of ox-ß2m at pH 8.0 as zero and that of ox-ß2m at pH 1.0 as unity and scaling all other values accordingly.
In experiments to measure the initial rate of fibrillogenesis, red-ß2m (0.2 mg/ml in 25 mM Tris HCl, 25 mM sodium acetate, 25 mM glycine, 25 mM MES) was prepared as described above and thio-T (10 µM final concentration) was added. Fibrillogenesis was then initiated by the addition of 0.4 M NaCl at 10 °C. The fluorescence of thio-T was then measured continuously (using an excitation wavelength of 444 nm and emission at 480 nm). The initial rate of fibrillogenesis was determined from the initial change in thio-T fluorescence with time observed over the first 20 sec of the experiment. The initial rate of growth was reproducible to within 10% over several experiments.
Electron microscopy
Colloidon coated copper EM grids were placed coated side down onto sample drops containing preformed ß2m fibrils for 20 sec. The grids were then blotted with filter paper to remove excess solvent and the sample was stained with 4% (wt/vol) uranyl acetate for 20 sec. Grids were then blotted again and air-dried before analysis. All images were taken using a Philips CM10 electron microscope operating at 100 keV.
Circular dichroism
CD experiments were performed on a Jasco J715 CD spectropolarimeter. For measurements in the far UV (210–260 nm) the CD signal was recorded at 10°C in a 1-mm path-length cell using a protein concentration of 0.2 mg/ml. For measurement in the near UV (250–350 nm), CD spectra were recorded using a protein concentration of either 0.4 mg/ml (at pH 7) or 0.6 mg/ml (at pH 3.6 and 1.5) and a 1-cm path-length cuvette. Measurements were recorded with a 1 nm bandwidth, 1 nm resolution, interval speed of 50 nm/min and response times of 8 sec. Six accumulations were taken and the results were averaged. DTT absorbs light strongly below 210 nm. As a consequence, far UV CD spectra could not be acquired below 210 nm.
ANS binding
The fluorescence emission spectra of ANS with and without protein were recorded on a Perkin-Elmer LS50B spectrofluorimeter at 10°C. Protein samples (10 µM) were prepared as described as above and were diluted 10-fold into buffer containing 25 mM Tris HCl, 25 mM sodium acetate, 25 mM glycine, 25 mM MES and ANS (250 µM final concentration) at the appropriate pH value. ANS binding was determined immediately using an excitation wavelength of 389 nm and fluorescence emission was collected between 400 and 600 nm using slit widths of 5 nm. The fluorescence emission of ANS in buffer alone was then subtracted from that in the presence of ß2m.
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Acknowledgments |
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References |
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Ayers, D.C., Athanasou, N.A., Woods, C.G., and Duthie, R.B. 1993. Dialysis arthropathy of the hip. Clin. Orthopaed. Rel. Res. 290: 216–224.
Bellotti, V., Stoppini, M., Mangione, P., Sunde, M., Robinson, C., Asti, L., Brancaccio, D., and Ferri, G. 1998. Beta-2-microglobulin can be refolded into a native state from ex vivo amyloid fibrils. Eur. J. Biochem. 258: 61–67.[Medline]
Berggard, I. and Bearn, A.G. 1968. Isolation and properties of a low molecular weight ß2-globulin occurring in human biological fluids. J. Biol. Chem. 243: 4095–4103.
Bjorkman, P.J., Saper, M.A., Samaoui, B., Bennett, W.S., Strominger, J.L., and Wiley, D.C. 1987. Structure of the human class I histocompatibity antigen, HLA-A2. Nature 329: 506–512.[CrossRef][Medline]
Booth, D.R., Sunde, M., Bellotti, V., Robinson, C.V., Hutchinson, W.L., Fraser, P.E., Hawkins, P.N., Dobson, C.M., Radford, S.E., Blake, C.C.F., and Pepys M.B. 1997. Instability, unfolding and aggregation of human lysozyme variants underlying amyloid fibrillogenesis. Nature 385: 787–793.[CrossRef][Medline]
Bouchard, M., Zurdo, J., Nettleton, E.J., Dobson, C.M., and Robinson, C.V. 2000. Formation of insulin amyloid fibrils followed by FTIR simultaneosly with CD and electron microscopy. Protein Sci. 9: 1960–1967.[Abstract]
Bowden, N.B., Weck, M., Choi, I.S., and Whitesides, G.M. 2001. Molecule-mimetic chemistry and mesoscale self-assembly. Acc. Che. Res. 34: 231–238.[CrossRef][Medline]
Brancaccio, D., Ghiggeri, G.M., Braidotti, P., Garberi, A., Gallieni, M., Bellotti, V., Zoni, U., Gusmano, R., and Coggi, G. 1995. Deposition of kappa-light-chains and lambda-light-chains in amyloid filaments of dialysis-related amyloidosis. J. Am. Soc. Neph. 6: 1262–1270.
Campistol, J.M., Bernard, D., Papastoitsis, G., Sole, M., Kasirsky, J., and Skinner, M. 1996. Polymerization of normal and intact beta 2-microglobulin as the amyloidogenic protein in dialysis-amyloidosis. Kidney Int. 50: 1262–1267.[Medline]
Chiti, F., Webster, P., Taddei, N., Clark, A., Stefani, M., Ramponi, G., and Dobson, C.M. 1999. Designing conditions for in vitro formation of amyloid protofilaments and fibrils. Proc. Natl. Acad. Sci. 96: 3590–3594.
Colfen, H., Harding, S.E., Wilson, E.K., Packman, L.C., and Scrutton, N.S. 1996. Homodimeric and expanded behavior of trimethylamine dehydrogenase in solution at different temperatures. Eur. Biophys. J. 24: 159–164.
Connors, L.H., Shirahama, T., Skinner, M., Fenves, A., and Cohen, A.S. 1985. in vitro formation of amyloid fibrils from intact beta 2-microglobulin. Biochem. Biophys. Res. Comm. 131: 1063–1068.[CrossRef][Medline]
Esposito, G., Michelutti, R., Verdone, G., Viglino, P., Herandez, H., Robinson, C.V., Amoresano, A., Dal Plaz, F., Monti, M., Pucci, P., et al. 2000. Removal of the N-terminal hexapeptide from human beta-2-microglobulin facilitates protein aggregation and fibril formation. Protein Sci. 9: 831–845.[Abstract]
Floege, J. and Ehlerding, G. 1996. Beta 2-microglobulin associated amyloidosis. Clin. Issues in Nephrol. 72: 9–26.
Frisch, C., Kolmar, H., Schmidt, A., Kleemann, G., Reinhardt, A., Pohl, E., Uson, I., Schneider, T.R., and Fritz, H.J. 1996. Contribution of the intramolecular disulfide bridge to the folding stability of REI(v), the variable domain of a human immunoglobulin kappa light chain. Folding & Design 1: 431–440.[CrossRef][Medline]
Gejyo, F., Homma, N., Suzuki, Y., and Arakawa, M. 1986. Serum levels of beta-2-microglobulin as a new form of amyloid protein in patients undergoing long-term hemodialysis. N. Engl. J. Med. 314: 585–586.[Medline]
Gejyo, F., Kazama, J.J., Hasegawa, S., Nishi, S., Arakawa, M., and Odano, I. 1995. 131I-beta-2-microglobulin scintigraphy in patients with dialysis amyloidosis. Clin. Nephrol. 44 (Suppl 1): S14–S18.
Glenner, G.G. 1980. Amyloid deposits and amyloidosis. The beta-fibrilloses. N. Engl. J. Med. 302: 1283–1292.[Medline]
Goldsbury, C.S., Cooper, G.J.S., Goldie, K.N., Muller, S.A., Saafi, E.L., Gruijters, W.T.M., and Misur, M.P. 1997. Polymorphic fibrillar assembly of human amylin. J. Struct. Biol. 119: 17–27.[CrossRef][Medline]
Guex, N. and Peitsch, M.C. 1997. Swiss-Model and the Swiss-Pdb Viewer: An enviroment for comparative protien modeling. Electrophoresis 18: 2714–2723.[CrossRef][Medline]
Guijarro, J.I., Sunde, M., Jones, J.A., Campbell, I.D., and Dobson, C.M. 1998. Amyloid fibril formation by an SH3 domain. Proc. Natl. Acad. Sci. 95: 4224–4228.
Helms, L.R., and Wetzel, R. 1996. Specificity of abnormal assembly in immunoglobulin light chain deposition disease and amyloidosis. J. Mol. Biol. 257:77–86.[CrossRef][Medline]
Hou, F.F., Miyata, T., Boyce, J., Yuan, Q., Chertow, G.M., Kay, J., Schmidt, A.M., and Owen, W.F. 2001a. Beta-2-microglobulin modified with advanced glycation end products delays monocyte apoptosis. Kindey Int. 59: 990–1002.[CrossRef][Medline]
Hou, F.F., Reddan, D.N., Seng, W.K., and Owen, W.F. 2001b. Pathogenesis of beta-2-microglobulin amyloidosis: Role of monocytes/macrophages. Semin. Dial. 14: 135–139.[CrossRef][Medline]
Inoue, S., Kuroiwa, M., Ohashi, K., Hara, M., and Kisilevsky, R. 1997. Ultrastructural organization of hemodialysis-associated beta-2-microglobulin amyloid fibrils. Kidney Int. 52: 1543–1549.[Medline]
Isenman, D.E., Painter, R.H., and Dorrington, K.J. 1975. The structure and function of immunoglobulin domains: Studies with beta-2-microglobulin on the role of the intrachain disulfide bond. Proc. Natl. Acad. Sci. 72: 548–552.
Jimenez, J.L., Guijarro, J.L., Orlova, E., Zurdo, J., Dobson, C.M., Sunde, M., and Saibil, H.R. 1999. Cryo-electron microscopy structure of an SH3 amyloid fibril and model of the molecular packing. EMBO J. 18: 815–821.[CrossRef][Medline]
Klafki, H.W., Pick, A.I., Pardowitz, I., Cole, T., Awni, L.A., Barnikol, H.U., Mayer, F., Kratzin, H.D., and Hilschmann, N. 1993. Reduction of disulphide bonds in an amyloidogenic Bence Jones protein leads to formation of amyloid-like fibrils in-vitro. Biol. Chem. Hoppe-Seyler 374: 1117–1122.[Medline]
Klunk, W.E., Jacob, R.F., and Mason, R.P. 1999. Quantifying amyloid beta-peptide (Ab) aggregation using the Congo red Ab (CR-Ab) spectrophotometric assay. Anal. Biochem. 266: 66–76.[CrossRef][Medline]
Kocisko, D.A., Lansbury, P.T., and Caughey, B. 1996. Partial unfolding and refolding of scrapie-associated prion protein: Evidence for a critical 16-kDa C-terminal domain. Biochemistry 35: 13434–13442.[CrossRef][Medline]
Lai, Z.H., Colon, W., and Kelly, J.W. 1996. The acid-mediated denaturation pathway of transthyretin yields a conformational intermediate that can self-assemble into amyloid. Biochemistry 35: 6470–6482.[CrossRef][Medline]
Lansbury, P.T. Jr. 1999. Evolution of amyloid: What normal protein folding may tell us about fibrillogenesis and disease. Proc. Natl. Acad. Sci. 96: 3342–3344.
McParland, V.J., Kad, N.M., Kalverda, A.P., Brown, A., Kirwin-Jones, P., Hunter, M.G., Sunde, M., and Radford, S.E. 2000. Partially unfolded states of beta-2-microglobulin and amyloid formation in vitro. Biochemistry 39: 8735–8746.[CrossRef][Medline]
Morgan, C.J., Gelfand, M., Ateya, C., and Miranker, A.D. 2001. Kidney dialysis-associated amyloidosis: A molecular role for copper in fiber formation. J. Mol. Biol. 309: 339–345.[CrossRef][Medline]
Naiki, H., Higuchi, K., Hosokawa, M., and Takeda, T. 1989. Fluorometric-determination of amyloid fibrils in vitro using the fluorescent dye, Thioflavin-T. Anal. Biochem. 177: 244–249.[CrossRef][Medline]
Naiki, H., Hashimoto, N., Suzuki, S., Kimura, H., Nakakuki, K., and Gejyo, F. 1997. Establishment of a kinetic model of dialysis-related amyloid fibril extension in vitro. Amyloid-Inter. J. Exp. Clin. Inv. 4: 223–232.
Nishi, S., Ogino, S., Maruyama, Y., Honma, N., Gejyo, F., Morita, T., and Arakawa, M. 1990. Electron-microscopic and immunohistochemical study of beta-2-microglobulin-related amyloidosis. Nephron 56: 357–363.[Medline]
Ohashi, K., Hara, M., Kawai, R., Ogura, Y., Honda, K., Nihei, H., and Mimura, N. 1992. Cervical disks are most susceptible to beta-2-microglobulin amyloid deposition in the vertebral column. Kidney Int. 41: 1646–1652.[Medline]
Okon, M., Bray, P., and Vucelic, D. 1992. 1H-NMR assignments and secondary structure of human beta-2-microglobulin in solution. Biochemistry 31: 8906–8915.[CrossRef][Medline]
Ono, K. and Uchino, F. 1994. Formation of amyloid-like substance from beta-2-microglobulin in vitro. Nephron 66: 404–407.[Medline]
Perfetti, V., Ubbiali, P., and Vignarelli, M.C. 1998. Evidence that amyloidogenic light chains undergo antigen-driven selection. Blood 91: 2948–2954.
Raffen, R., Dieckman, L.J., Szpunar, M., Wunschl, C., Pokkuluri, P.R., Dave, P., Stevens, P.W., Cai, X.Y., Schiffer, M., and Stevens, F.J. 1999. Physicochemical consequences of amino acid variations that contribute to fibril formation by immunoglobulin light chains. Protein Sci. 8: 509–517.[Abstract]
Ritz, E. and Zeier, M. 1996. Beta-2-microglobulin associated amyloidosis. Clin. Issues Nephrol. 72: 9–26.
Stevens, F.J., Pokkuluri, P.R., and Schiffer, M. 2000. Protein conformations and disease: Pathological consequences of analogous mutations in homologous proteins. Biochemistry 39: 15291–15296.[CrossRef][Medline]
Sunde, M. and Blake, C. 1997. The structure of amyloid fibrils by electron microscopy and X-ray diffraction. Adv. Protein Chem. 50: 123–159.[Medline]
Sunde, M., Serpell, L.C., Bartlam, M., Fraser, P.E., Pepys, M.B., and Blake, C.C.F. 1997. Common core structure of amyloid fibrils by synchrotron X-ray diffraction. J. Mol. Biol. 273: 729–739.[CrossRef][Medline]
Swietnicki, W., Petersen, R., Gambetti, P., and Surewicz, W.K. 1997. pH-dependent stability and conformation of the recombinant human prion protein PrP(90–231). J. Biol. Chem. 272: 27517–27520.
Swietnicki, W., Morillas, M., Chen, S.G., Gambetti, P., and Surewicz, W.K. 2000. Aggregation and fibrillisation of the recombinant human prion protein huPrP90–231. Biochemistry 39: 424–431.[CrossRef][Medline]
Van Ypersele, C. and Drucke, T.B. 1996. Chapter two: Various clinical types of amyloidosis. In Dialysis Amyloid. pp. 34–68. Oxford University Press, UK.
Whitesides, G.M., Mathias, J.P., and Seto, C.T. 1991. Molecular self-assembly and nanochemistry: A chemical strategy for the synthesis of nanostructures. Science 254: 1312–1319.
Woody, R.W. 1995. Circular-dichroism. Methods in Enzymol. 246: 34–71.[Medline]
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) Red-ß2m; (•) ox-ß2m. All spectra were acquired at 10°C and a protein concentration of 0.2 mg/ml. Far UV CD spectra at (A) pH 7.0, (B) pH 3.6, (C) pH 1.5. Near UV CD spectra at (D) pH 7.0, (E) pH 3.6 and (F) pH 1.5.
) ANS alone at (A) pH 7.0, (B) pH 3.6, and (C) pH 1.5. All spectra were acquired at 10°C.
) red-ß2m in the presence of 0.4 M NaCl; and (
