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. 2005 Jul;14(7):1811–1817. doi: 10.1110/ps.051387005

Urea-induced denaturation of apolipoprotein serum amyloid A reveals marginal stability of hexamer

Limin Wang 1,1, Wilfredo Colón 1
PMCID: PMC2253367  PMID: 15937280

Abstract

Serum Amyloid A (SAA) is an acute phase reactant protein that is predominantly found bound to high-density lipoprotein in plasma. Upon inflammation, the plasma concentration of SAA can increase dramatically, occasionally leading to the development of amyloid A (AA) amyloidosis, which involves the deposition of SAA amyloid fibrils in major organs. We previously found that the murine isoform SAA2.2 exists in aqueous solution as a hexamer containing a central channel. Here we show using various biophysical and biochemical techniques that the SAA2.2 hexamer can be totally dissociated into monomer by ~2 M urea, with the concerted loss of its α-helical structure. However, limited trypsin proteolysis experiments in urea showed a conserved digestion profile, suggesting the preservation of major backbone topological features in the urea-denatured state of SAA2.2. The marginal stability of hexameric SAA2.2 and the presence of residual structure in the denatured monomeric protein suggest that both forms may interconvert in vivo to exert different functions to meet the various needs during normal physiological conditions and in response to inflammatory stimuli.

Keywords: amyloidosis, HDL, inflammation, acute phase, amyloid fibril

Serum Amyloid A (SAA) belongs to a family of proteins that play a major role in the acute phase inflammatory response (Uhlar and Whitehead 1999). As a result of inflammation, the concentration of SAA in plasma can increase up to 1000 times (McAdam and Sipe 1976). In vivo, SAA predominantly associates with the third fraction of high-density lipoprotein (HDL) (Benditt and Eriksen 1977). To date, SAA has been found to exist with a high degree of sequence homology in all vertebrate species examined, indicating that SAA is a highly conserved protein (Uhlar et al. 1994). The high degree of conservation, the wide expression profile in various tissues/ cells besides the liver, and the dramatic increase in expression levels during inflammation, suggest that SAA may play important beneficial roles in host defense. However, persistently high expression of SAA during chronic inflammatory disease may result in AA amyloidosis (formerly reactive amyloidosis), which is characterized by the deposition of amyloid fibrils usually involving the N-terminal 76-residue fragment (amyloid A) in major organs like the liver, the spleen, and the kidney (Husebekk et al. 1985; Gillmore et al. 2001). Consistent with the evidence that atherosclerosis is an inflammatory disease (Libby et al. 2002), SAA’s serum concentration has been proposed to be a better prediction marker of coronary heart disease than cholesterol (Delanghe et al. 2002; Saadeddin et al. 2002).

Although SAA appears to have very important functions in vivo, these remain largely obscure, because the high tendency of HDL-free SAA to aggregate in vitro has limited experimental work, including the solution of its three-dimensional structure. Recently, we found that murine SAA2.2, a nonamyloidogenic isoform in vivo, has modest solubility and forms a hexameric structure with a putative central channel (Wang et al. 2002). To investigate the stability of SAA2.2, we used various biochemical and biophysical techniques to monitor its structural changes upon urea-induced denaturation. Here we show that hexameric SAA2.2 is marginally stable and undergoes a concerted loss of quaternary, tertiary, and secondary structure. These results have implications concerning the in vivo amyloidogenicity and the multiple functions of SAA.

Results

Hexamer to monomer denaturation without oligomeric intermediates

The quaternary structure of SAA2.2 can be conveniently trapped using glutaraldehyde cross-linking (GCL), which basically involves the covalent reaction between the reagent’s two aldehyde groups and the lysine residue’s ɛ-amino group (Craig 1988). When SAA2.2 was incubated in different concentrations of urea, GCL clearly showed a hexamer to monomer transition without population of dimeric or trimeric intermediates (Fig. 1 ▶). Interestingly, hexamer dissociation began at the lowest concentration of urea (0.1 M), and there was nearly complete dissociation at 2.1 M urea.

Figure 1.

Figure 1.

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SAA2.2 urea-induced hexamer to monomer dissociation monitored by glutaraldehyde cross-linking and SDS-PAGE. SAA2.2 samples contained 50 μg/mL of protein in MOPS buffer at pH 7.4 and were preincubated at 20°C for 10 min before cross-linking with 0.7% (v/v) glutaraldehyde.

To directly monitor the quaternary structure of SAA2.2, we also used size-exclusion chromatography (SEC). In the SEC experiment, the peak corresponding to hexameric SAA2.2 begins to decrease at low urea concentrations (0.5 M) and disappears by 2 M urea (Fig. 2A ▶), as in the GCL experiment (Fig. 1 ▶). Concomitant with the disappearance of the hexameric peak, another peak with greater retention time, presumably corresponding to the monomer, increases. The retention time of this peak decreases as the urea concentration increases up to 6 M (Fig. 2B ▶), perhaps due to a ureainduced increase in the hydrodynamic radius of the denatured protein. The dramatic effect of urea on the retention time of the denatured monomeric SAA2.2 suggests that the hexamer dissociates to a relatively compact denatured monomeric state.

Figure 2.

Figure 2.

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SAA2.2 urea-induced quaternary structural changes monitored by size exclusion chromatography. (A) Hexamer to monomer transition is completed by 2 M urea. (B) At urea concentrations higher than 2 M, there is a decrease in the monomer peak retention time. SAA2.2 samples contained 0.1 mg/mL protein in 20 mM Tris buffer with 0.4M NaCl. Separation was achieved using a Superdex 75 PC 3.2/30 column at a flow rate of 0.1 mL/min.

Structural changes monitored by fluorescence polarization

SAA2.2 has three tryptophan (Trp) residues at positions 17, 28, and 52, and therefore, we used them as probes to determine the fluorescence polarization as a function of urea concentration to obtain more information about the quaternary (and probably tertiary) changes of SAA2.2. The polarization data (Fig. 3A ▶) is quite consistent with the results from the GCL and SEC experiments, suggesting that SAA2.2 dissociation from hexamer to monomer is the major factor in the decrease in fluorescence polarization. The polarization of SAA2.2 changed with increasing urea concentration up to 3 M urea, after which it remained constant. The fluorescence spectra showed a red shift in the wavelength of maximum fluorescence emission (λmax) from ~340 nm (without urea) to ~354 nm (at 5.3 M urea) (Fig. 3B ▶), indicating complete solvent exposure of the Trp residues (Colón 1999). Interestingly, there was little change in λmax in the 0–2 M urea range, suggesting that the hexamer dissociated to a monomeric structure with one or more Trp residues partially protected from the solvent.

Figure 3.

Figure 3.

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SAA2.2 urea-induced denaturation monitored by fluorescence polarization. (A) Fluorescence polarization plotted as a function of urea concentration. SAA2.2 samples contained 50 μg/mL of protein in MOPS buffer at pH 7.4 and were incubated at 20°C for 10 min. The symbols represent three independent experiments. (B) Fluorescence emission spectra of SAA2.2 samples containing the indicated urea concentrations.

Loss of α-helical structure

Far-UV circular dichroism (CD) was performed to examine the secondary structure changes of SAA2.2 upon urea denaturation. Since native hexameric SAA2.2 possesses ~50% α-helical and ~10% β-sheet structure (Wang et al. 2002), the CD signal is mainly monitoring the presence of α-helical structure. The CD spectra recorded at various urea concentrations show a dramatic loss of α-helical content at low urea concentrations (Fig. 4A ▶). When the signal was monitored at 222 nm, the resulting transition lacked a pretransition baseline and showed that most of the native structure is lost by ~2Murea (Fig. 4B ▶), similar to what was seen by the other methods (Figs. 1 ▶–3 ▶ ▶).

Figure 4.

Figure 4.

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SAA2.2 urea denaturation monitored by far UV CD. SAA2.2 samples contained 50 μg/mL of protein in MOPS buffer at pH 7.4 and were incubated at 20°C for 10 min. (A) Wavelength scans taken at some urea concentrations of the sample. (B) Molar residue ellipticity [θ] at 222 nm plotted against urea concentration. Symbols represent two independent experiments.

Limited trypsin proteolysis suggest conserved topological features

The protease trypsin, which cleaves proteins after Arg and Lys residues, was used to determine the changes in accessible proteolytic sites as a function of urea. There are 13 potential trypsin cleavage sites at positions 18, 24, 29, 33, 38, 46, 56, 61, 70, 83, 86, 89, and 102 along the 103-residue SAA2.2 protein (Wang et al. 2002). Therefore, limited trypsin proteolysis may reveal which SAA2.2 regions are more solvent exposed and susceptible to proteolysis at increasing concentrations of urea. SAA2.2 samples in the absence or the presence of urea were incubated with trypsin for a certain amount of time before quenching the reaction with trifluoroacetic acid, and the resulting reverse-phase HPLC analysis surprisingly showed similar chromatograms for all of the SAA2.2 samples (Fig. 5 ▶). Among the potential digestion sites, Arg 38 and Arg 86 were always the two that were preferentially cleaved to yield the 39–86 SAA2.2 fragment. The identity of the fragments generated by limited trypsin proteolysis of SAA2.2 in the absence of urea was previously determined by mass spectrometry (Wang et al. 2002). These results suggest that the denatured monomeric SAA2.2 in urea has some residual structure that retains Arg 38 and Arg 86 as the most accessible cleavage sites.

Figure 5.

Figure 5.

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Limited trypsin digestion and reverse-phase HPLC analysis of SAA2.2 samples incubated in different concentrations of urea. SAA2.2 samples contained 0.1 mg/mL protein in MOPS buffer at pH 7.4 and were preincubated at 20°C for 30 min before limited trypsin digestion at an SAA:trypsin ratio of 120:1. The reaction was quenched with trifluoroacetic acid and analyzed by reverse-phase HPLC as described in Materials and Methods.

Discussion

Denaturation of SAA2.2 involves the concerted loss of quaternary, tertiary, and secondary structure

The denaturation mechanism of SAA2.2 was investigated using several biochemical and biophysical methods (GCL, SEC, fluorescence polarization, and CD) to monitor different aspects of its structure. The denaturation data was similar regardless of the method used, as indicated by the nearly superimposable transitions, with the exception of the polarization data, which is slightly shifted to the right (Fig. 6 ▶). This inconsistency is not uncommon when using intrinsic chromophores to measure polarization, and could be caused by several factors, like differences in fluorophore lifetime between the native and denatured states, contribution of monomer unfolding to the polarization, or urea-induced changes in solution viscosity (Eftink 1994; Lakowicz 1999). Nevertheless, the similar urea-induced denaturation transitions of SAA2.2 when monitored by GCL, SEC, and CD, suggest that the denaturation mechanism of SAA2.2 predominantly involves the concerted loss of quaternary, tertiary, and secondary structure, thereby suggesting that the oligomerization of SAA2.2 may play a major role in stabilizing its secondary and tertiary structure. Therefore, the urea-induced denaturation of SAA2.2 should exhibit a strong protein concentration dependence. Unfortunately, we were not able to successfully carry out these experiments, due to aggregation of SAA2.2 at higher concentrations and decreased signal-to-noise ratio at lower SAA2.2 concentrations. We propose that the N terminus of SAA2.2 is involved in the oligomerization of SAA2.2, based on the observation that only monomeric SAA2.2 can bind to HDL in vitro (Wang and Colón 2004), thereby suggesting that the HDL-binding N terminus of SAA2.2 (Liang et al. 1996; Patel et al. 1996) is not available for binding within the hexamer. Furthermore, the SAA2.2 central fragment involving residues 39–86 does not hexamerize, and the C terminus of SAA2.2 is a proline-rich region predicted to be disordered (Wang et al. 2002).

Figure 6.

Figure 6.

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Comparison of the urea denaturation curves obtained from different methods. The GCL/SDS–polyacrylamide gel was analyzed, and the monomer percentage was plotted against urea concentration. For the SEC curve, the fractional area of the slow-moving SAA2.2 peak in SEC was used. The polarization and CD data was internally normalized using the values at 0 (0%) and 5.8M (100%) urea.

The observation of a conserved initial trypsin proteolytic profile in the presence of urea suggests that the denatured monomeric SAA2.2 retains some of the topological features present in the hexamer. This is consistent with the results showing a further red shift in the fluorescence maximum wavelength (Fig. 3B ▶) upon increasing the urea concentration from 2 to 5 M. Similarly, the monomer eluted from the size-exclusion HPLC column at shorter retention times with increasing urea concentrations from 2–5 M, indicating further unfolding and expansion of the monomer. Thus, urea denatures hexameric SAA2.2 into a monomeric denatured state, lacking most of its native secondary and tertiary structure, but retaining some of its main chain topology.

Hexameric SAA2.2 is marginally stable

The changes in quaternary (Figs. 1 ▶–3 ▶ ▶) and secondary (Fig. 4 ▶) structures of SAA2.2 at urea concentrations as low as 0.1 M clearly show that hexameric SAA2.2 is a marginally stable protein. Due to the lack of a pretransition baseline, it was not possible to fit the data to obtain a reliable free-energy change of unfolding. Nevertheless, the clear qualitative observation of a broad denaturation transition (i.e., low m-value) that ranges from 0 to 2–3 M urea, combined with the low-denaturation mid-point (cm), indicate that hexameric SAA2.2 has marginal stability. Often, the very low in vitro stability exhibited by some proteins is due to the absence of ligands or cofactors, which play a stabilizing role. In addition to its ability to bind HDL, it has been shown or proposed that SAA may bind a number of ligands in vivo, including calcium (Turnell et al. 1986), integrin and fibronectin-like binding elements (Preciado-Patt et al. 1994), and heparin/heparan sulfate (Ancsin and Kisilevsky 1999a). Thus, it is possible that hexameric SAA2.2 may be stabilized in vivo by binding to these or other ligands. It is also worth noting that during inflammation, the SAA concentration may increase (up to 1000-fold) to above 1 mg/mL, which is 10–20 times higher than the concentrations used in this study. Such high SAA concentration could significantly stabilize the hexamer.

The marginal stability of SAA2.2 shown here is in agreement with our observation that even at physiological temperature (37°C), SAA2.2 dissociates into a monomer and loses most of its secondary structure (Wang et al. 2005). As shown in the present study, the temperature-induced denaturation of SAA2.2 involved a hexamer to monomer transition without significant population of intermediates, and even though most of the secondary structure of SAA2.2 was lost at 37°C, limited proteolysis experiments also showed that the denatured monomer retained some of the topological features of the hexamer. Therefore, it seems that at 37°C, monomeric SAA has structural properties similar to “natively unfolded” proteins, a class of proteins that are mainly characterized by lacking well-defined structure in vitro (Wright and Dyson 1999; Uversky 2002). Thus, unless the hexameric protein is stabilized in vivo by ligand binding or by the dramatic increase in its concentration as a result of inflammation, SAA may exist as a monomeric natively unfolded protein. This may explain in part SAA’s ability to bind many ligands, as it is becoming accepted that the natively unfolded structure of many proteins may be required for molecular recognition functions, such as oligomerization and binding to many partners (Dafforn and Smith 2004).

Implications for SAA function and amyloid fibril formation in vivo

The marginal stability of SAA is likely to have in vivo functional consequences. For example, the hexameric and monomeric forms of SAA may bind to different ligands, as suggested by the observation that monomeric, but not hexameric, SAA2.2 binds to HDL in vitro (Wang and Colón 2004). Therefore, the marginal stability of SAA may modulate its functions by allowing easy conversion between different oligomeric forms. In this context, the increase in SAA concentration during inflammation may stabilize the hexamer, and thereby serve as a regulatory mechanism to activate host defense functions required during an acute phase response. In addition, the ability of monomeric SAA2.2 to retain some of the hexamer topological features when denatured under mild denaturing conditions (2 M urea or 37°C) in vitro suggests that it may have other in vivo functions aside from HDL binding. This would help explain the large number of functions related to cholesterol metabolism and innate immunity that have been attributed to SAA (Uhlar and Whitehead 1999).

The marginal stability of SAA2.2 may also have a profound impact on the ability of amyloidogenic SAA isoforms to deposit into amyloid fibrils in vivo, leading to AA amyloidosis. Preliminary data have shown that the murine SAA1.1 isoform, which is amyloidogenic in vivo, is also a very unstable protein (L. Wang and W. Colón, unpubl.). The very high (97/103) sequence identity between the SAA2.2 and the amyloidogenic SAA1.1 isoforms, together with our observation that SAA2.2 can easily form amyloid fibrils in vitro at 37°C (Wang et al. 2005), suggest that although SAA2.2 is nonamyloidogenic in vivo, it still possess some of the intrinsic amyloidogenicity of SAA1.1. Thus, SAA2.2 may be a good model for understanding the mechanism of amyloid formation by SAA1.1. Since the formation of SAA amyloid fibrils involves the misfolding into a β-rich amyloidogenic species, the marginal stability of SAA is expected to facilitate its misfolding. This is consistent with the observation that at 37°C, SAA2.2 dissociates in vitro and misfolds into a β-rich monomeric species that self-assembles into amyloid fibrils (Wang et al. 2005). Although the marginal stability of SAA may increase its susceptibility to amyloid formation, it may also provide a means for allowing the rapid turnover of the protein. This may be especially important in the case of SAA, since its concentration can increase up to 1000 times in 24 h upon an inflammatory insult (McAdam and Sipe 1976). Since the occurrence of AA amyloidosis is relatively rare among patients suffering from chronic inflammatory diseases, it appears that other in vivo factors, such as ligand binding and an efficient quality control system, play an important role in keeping this marginally stable protein from frequently depositing into amyloid fibrils. In similar fashion, a compromised quality control system and the abnormal interaction of SAA with ligands or membranes may enhance its ability to deposit into amyloid fibrils (Ancsin and Kisilevsky 1999b). Thus, a better understanding of the mechanism of SAA amyloid formation in vivo and the role of SAA in other pathological processes (Urieli-Shoval et al. 2000) will require elucidating the relationship amongst its structure, stability, and ligand binding, and understanding how these are modulated in vivo.

Materials and methods

SAA expression and purification

The murine SAA2.2 cDNA was cloned into a pET21-a(+) vector between the NdeI and BamHI sites and transformed into Escherichia coli strain BL21 (DE3) pLysS-competent cells, as previously described (Liang et al. 1998). The expression and purification procedure was modified from that of Yamada et al. (1994) and is described in detail in our previous publications (Wang et al. 2002; Wang and Colón 2004). Basically, the IPTG-induced cells were lysed by three cycles of freezing/thawing and then incubated in 2-amino-2-hydroxy-methyl- 1,3-propanediol buffer (Tris, 20 mM [pH 8.2]) containing 6 M urea. The soluble extract was loaded onto a DEAE anion exchange column, and the fractions containing SAA2.2 were pooled and loaded onto a chromatofocusing column. The tubes containing pure SAA2.2 were precipitated with ammonium sulfate (70% saturation), and the centrifuged pellet was dissolved in Tris buffer or 20 mM 3-(N-morpholino) propane-sulfonic acid buffer (MOPS) at pH 7.4.

Glutaraldehyde cross-linking

SAA2.2 samples (50 μg/mL in MOPS buffer [pH 7.4], in the absence or the presence of urea) were preincubated at 20°C for 10 min, and then cross-linked with 0.7% (v/v) glutaraldehyde for 20 min as previously described (Wang et al. 2002). The crosslinking reaction was quenched by the addition of Tris and the extent of cross-linking was analyzed by SDS-PAGE. The gel was digitally photographed and quantitatively analyzed using the AlphaEaseFC software (Alpha Innotech Corp.).

Reverse phase and size-exclusion chromatography

A Gold Noveau Beckman Coulter HPLC instrument was used with a single wavelength absorbance detector (set at 220 nm). For analysis of limited trypsin digestion products, an analytic 4.6 mm×25 cm C4 reverse phase column (Vydac) was used at a flow rate of 0.66 mL/min. A 5%–90% linear gradient of aqueous solution (0.1% [v/v] trifluoroacetic acid and 90% [v/v] acetonitrile) was applied over 85 min, and the column was regenerated with 0.1% trifluoroacetic acid.

To monitor the urea-induced quaternary structural change, SAA2.2 samples (20 μL of 0.1 mg/mL, without or with urea) were incubated at room temperature for 10 min, and then analyzed by size exclusion chromatography on a Superdex 75 PC 3.2/30 column (Amersham Pharmacia Biotech) at a flow rate of 0.1 mL/min. The elution buffer (20 mM Tris buffer with 0.4 M NaCl) was the same as the sample buffer, including the same concentration of urea.

Fluorescence polarization spectroscopy

When a fluorescent molecule is excited by plane-polarized light, it will emit plane-polarized light if the molecule remains stationary between the time of excitation and emission (Lakowicz 1999). Therefore, the degree of polarization depends on the molecule’s rotational correlation time. The faster the molecule rotates and tumbles, i.e., the shorter the rotational correlation time, the smaller the polarization value will be. Since the rotational correlation time is based on the size and volume of the protein, fluorescence polarization can be used to study the quaternary structure changes in protein denaturation studies (Checovich et al. 1995). The polarization value is calculated as (I00-00−G*I00-90)/ (I00-00+G*I00-90), while G=I90-00/I90-90. The first two digits in the fluorescence intensity (I) subscript represent the rotation in degrees (0° or 90°) of the polarizer lens in the excitation side, while the last two digits are for the polarizer on the emission side. The factor G accounts for the transmission difference of the two polarizers.

A Hitachi F-4500 fluorescence spectrophotometer was used with excitation wavelength at 295 nm and a slit width of 5 nm. Excitation at 295 nm was used to avoid exciting the six Tyr residues present in SAA2.2. The emission slit width was set at 10 nm. A polarizer lens was installed at the excitation as well as the emission side and arranged at either 0° or 90° rotation from each other. Two stock samples (50 μg/mL SAA2.2 in MOPS buffer) were made with no urea or containing 9 M urea. Individual samples were prepared by a codilution method, in which SAA2.2 samples of progressively higher urea concentrations were made by repetitive withdrawal from a SAA2.2 sample (originally in 0 M urea) in the cell, followed by addition of the same volume of the SAA2.2 stock sample in 9 M urea. At each urea concentration, the sample was incubated for 10 min at 20°C before the 20-sec time scan. The signal did not change with longer incubation time, indicating that equilibrium had been reached. The codilution method described here for making protein samples containing different concentrations of urea is only appropriate with proteins that denature reversibly. We have previously shown that hexameric SAA2.2 can be reversibly denatured after incubation in 6 M urea (Wang et al. 2002). This is consistent with our purification protocol, which is carried out under denaturing conditions.

Circular dichroism

Spectra were recorded on an OLIS CD instrument with a 0.1-cm pathway sample cuvette. The SAA2.2 samples with different concentrations of urea were made by the codilution method described above. Each sample was incubated at 20°C for 20 min before three wavelength scans were collected, with a wavelength increment of 0.5 nm and a time table of 2 sec (260–230 nm) and 8 sec (230–200 nm). Additionally, 2-min time scans at 222 and 250 nm were also recorded to generate reliable transition curves. The signal at 250 nm was used as an internal control to correct for small fluctuations in the baseline.

Limited proteolysis

SAA2.2 sample (60 μL of 0.1 mg/mL without or with a certain concentration of urea) was preincubated at 20°C for 30 min and then partially digested by TPCK-treated trypsin (Sigma) with an SAA:trypsin ratio of 120:1. To balance the lower trypsin activity in higher urea concentration, proteolysis time was adjusted to obtain a similar degree of full-length SAA2.2 degradation. The proteolysis reaction was stopped by adding 0.1% (v/v) trifluoroacetic acid and then analyzed by analytic C4 reverse-phase HPLC as described above.

Acknowledgments

We thank Dr. Dmitri Zagorevski for Mass Spectrometry analyses and the Department of Chemistry for support of the Mass Spectrometry facility. This work was supported by grants from the American Heart Association and NIH (R01NS42915) to W.C.

Abbreviations

  • SAA, serum amyloid A

  • MOPS, 3-(N-morpholino) propanesulfonic acid

  • Tris, 2-amino-2-hydroxymethyl-1,3-propanediol

  • GCL, Glutaraldehyde cross-linking

  • SEC, size exclusion chromatography

  • CD, circular dichroism

  • RT, room temperature

  • λmax, wavelength of maximum fluorescence emission

  • MRE or θ, molar residue ellipticity

Article published online ahead of print. Article and publication date are at http://www.proteinscience.org/cgi/doi/10.1110/ps.051387005.

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