Molecular dynamics simulations of the unfolding of ß2-microglobulin and its variants

Buyong Ma1 and Ruth Nussinov1,2,3

1Basic Research Program, SAIC–Frederick, Inc., Laboratory of Experimental and Computational Biology, NCI-FCRDC, Frederick, MD 21702, USA and 2Sackler Institute of Molecular Medicine, Department of Human Genetics and Molecular Medicine, Sackler School of Medicine, Tel Aviv University, Tel Aviv 69978, Israel

3 To whom correspondence should be addressed. e-mail: ruthn{at}ncifcrf.gov


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
In this study, we examined the unfolding processes of native ß2-microglobulin and two related variants, one with an N-terminal hexapeptide deletion {Delta}N6 and another with Lys57–Asp58 cleavage, by high-temperature molecular dynamics simulations. Three simulation models were used, molecular dynamics (MD) simulations with explicit water solvation, MD simulations with the CHARMM EEF1 force field and Langevin dynamics with the CHARMM EEF1 force field. Our simulations reproduce many of the experimentally observed structural changes. The most striking agreement is in the ß-strands to {alpha}-helix transition. In our simulations, strands ß3, ß4 and ß5 consistently change to {alpha}-helix, whereas ß8 changes to an {alpha}-helix only briefly. Through comparisons of the conformational behavior of the native, the {Delta}N6 and the Lys-cut ß2-m, using the three simulation methods, we identified the consensus conformational changes that differentiate between the native ß2-m and its two variants. We found that the main effect of the removal of the N-terminal hexapeptide is to increase the separation between strands ß2 and ß6 and to facilitate the ß to {alpha} transition. On the other hand, the lysine cleavage only increases the flexibility of strand ß5 and does not affect the interactions between strands ß2 and ß6. These conformational changes may relate to polymerization tendencies of these variants.

Keywords: amyloid/ß2-microglobulin/ß2-microglobulin variant/ß-strand {alpha}-helix transition/molecular dynamics simulation/protein folding/protein unfolding


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
As the light chain of the major histocompatibility complex (MHC) class I antigens in humans and other vertebrates, ß2-microglobulin 2-m) has been found in a variety of physiological fluids as well as on the surfaces of nearly all cells (Floege and Ketteler, 2001Go). These antigens play central roles in two widely studied activities involving immune recognition by T cells: the rejection of foreign tissue grafts through direct recognition of foreign MHC antigens and the recognition of viral and other antigens in conjunction with self-MHC antigens. Sequence homology studies indicate that ß2-m belongs to a ‘superfamily’ of proteins related to the Ig constant domains. The three-dimensional structure (Becker and Reeke, 1985Go) of ß2-m is strikingly similar to the structures of Ig domains, with root mean square deviations (r.m.s.d.s) of <1.2 Å. The ß2-m chain is folded into a typical ‘ß-barrel’ configuration dominated by two antiparallel pleated sheets, comprised of five and three strands, respectively (Figure 1A). Following the ß-strand naming scheme of Esposito et al. (Esposito et al., 2000Go), the five-stranded sheet contains ß1 (residues 1–16), ß2 (17–31), ß4 (42–52), ß5 (53–58) and ß6 (59–73); the three-stranded sheet contains ß3 (32–41), ß7 (74–84) and ß8 (85–98). Strand ß4 runs between the two layers. There is a disulfide bond connecting the two antiparallel pleated sheets between ß2 (Cys25) and ß7 (Cys79). In this study we used the crystal structure from bovine ß2-m (Becker and Reeke, 1985Go), which has the same functional activity as and high sequence identity (76%) with human ß2-m.



View larger version (43K):
[in this window]
[in a new window]
 
Fig. 1. The initial structures of the studied systems: (A) Native ß2-microglobulin (PDB code 1bmg); (B) truncated {Delta}N6 ß2-microglobulin; (C) ß2-microglobulin with the Lys57–Asp58 bond cut off. The strands are ß1, 1–16; ß2, 17–31; ß3, 32–41; ß4,42–52; ß5, 53–58; ß6, 59–73; ß7, 74–84; and ß8, 85–98.

 
Amyloidosis of ß2-m is the origin of a serious disease in patients with longstanding uremia and hemodialysis treatment (Floege and Ketteler, 2001Go). ß2-m undergoes extracellular amyloid deposition and causes dialysis-related amyloidosis, a type of amyloidosis in which the fibrils have a selective propism for the skeletal-muscle system (Stoppini et al., 1997Go). The mechanism responsible for the formation of the ß2-m amyloid fibrils is still unclear. Experimental studies indicate that amyloid formation may be inhibited by binding anti-(ß2-m) monoclonal antibody (mAb) to residues 92–99 (strand ß8) of ß2-m, while the binding to residues 20–41 (strands ß2 and ß3) and 63–75 (strand ß6) have no effect on ß2-m fibrillogenesis (Stoppini et al., 1997Go). ß2-m variants show different amyloidosis behavior patterns. Reduction of the disulfide bond destabilizes the native protein. However, amyloid fibril formation increases only at low pH (1.5–4.0), but not at neutral pH (Smith and Radford, 2001Go). Esposito et al. (Esposito et al., 2000Go) removed the N-terminal hexapeptide ({Delta}N6) from the human ß2-m. They found that compared with the intact protein the truncated ß2-m was less stable and had a higher tendency to aggregate, forming amyloid fibrils. ß2-m can be refolded into a native state from ex vivo amyloid fibrils (Bellotti et al., 1998Go). Through the refolding experiments, it was found that a variant of ß2-m cleaved at Lys58 (Lys-cut) did not exist in an amyloid fibril form and that the cleaved species did not form amyloids (Bellotti et al., 2001Go; Heegaard et al., 2001Go).

The function and malfunction of a protein and its properties are determined by the native folded structure as well as by the distribution and redistribution of its conformational substates (Ma et al., 2001Go). The distribution of the conformational substates is strongly related to amyloidogenicity (Carrell and Lomas, 1997Go). For example, prion conformational diseases are caused by an aberrantly folded form of the prion protein, PrPSc, covalently indistinguishable but conformationally distinct from the normal form of PrPC (Harrison et al., 1997Go). By studying protein unfolding trajectories and the related conformational change, we may get an insight into the conformational ensemble which may already exist in solution under native conditions, although under such conditions the populations of these non-native conformers may be low. The population of non-native conformers is not sufficient for amyloid formation, as demonstrated by reduction of the disulfide bond of ß2-m at neutral pH without an increase in amyloid formation (Smith and Radford, 2001Go). Currently, neither experimental nor computational approaches can describe amyloid formation in atomic detail. Additional studies of the conformational behavior of related proteins should increase our understanding of this process.

In this study, we examined the unfolding processes of intact ß2-m and two related variants ({Delta}N6 and Lys-cut), by high-temperature molecular dynamics simulations. High- temperature unfolding simulations need to be closely monitored, since the sampled comformations do not necessarily relate to states accessible during real protein unfolding. Cross examinations of simulation results under different conditions, with related protein species and with available experimental data, are crucial. In these studies, three simulation models were used: molecular dynamics simulations with explicit water solvation, molecular dynamics (MD) simulations with the the CHARMM EEF1 force field and Langevin dynamics also with the CHARMM EFF1 force field. We focus on the mechanism of change of native ß-structures and the related conformational distributions, in order to understand events that may be involved in the transition from the native to the amyloid state. Through comparisons of the conformational behavior of the native, {Delta}N6 and the Lys-cut ß2-m, using the three simulation protocols, we identified consensus conformational changes that differentiate between the native ß2-m and its two variants. In our simulations, strands ß3, ß4 and ß5 consistently change to {alpha}-helices. ß8 only briefly changes to an {alpha}-helix (Table I). We found that the main effect of removing the N-terminal hexapeptide is the increase in the separation between strands ß2 and ß6, while the lysine cleavage does not affect the interactions between strands ß2 and ß6.


View this table:
[in this window]
[in a new window]
 
Table I. Summary of CHARMM EEF1 simulation conditions and characteristic new secondary structurea,b formations observed in the simulations
 

    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
The native polypeptide conformation was taken from the PDB [ß2-microglobulin, 1bmg (Bernstein et al., 1977Go)] without crystal waters. Hydrogen atoms, if missing in the deposited structure, were added using the biopolymer module in the INSIGHTII package (BIOSYM/MSI).

Molecular dynamics simulations with explicit water solvation were performed in the canonical ensemble (NVT) using the program Discover 2.98. For the shell water simulation, the system consisted of the protein solvated with a 20 Å shell of water molecules. To allow sufficient freedom for protein unfolding, the system was resolvated with more water molecules when the protein molecule expanded and more of its surface was exposed. Recently, Gilquin et al. (Gilquin et al., 2000Go) have shown that immersing a smaller solvation box into a larger one during the simulation gives similar results to those obtained from simulations using a larger solvation box already at the initial stage. For the simulations with periodic boundary conditions, the protein was solvated in a 60x60x60 Å3 cubic box with 6565 water molecules. The effective water density in the solvation system was 1.006 g/cm3. All atoms of the system were considered explicitly and their interactions were computed using the CFF91 force field (Maple et al., 1998Go). The potential energy functions include bond stretching, angle bending, torsion and out-of-plane angle deformation terms. It also contains cross-terms to describe the couplings between bond–bond, angle–angle, bond–angle, bond–torsion, torsion–angle and angle–angle–torsion. The time step in the MD simulations was 1 fs. Snapshots from the trajectories were saved every 1 ps. Only the trajectories of the protein were saved. Those of the water molecules were discarded to save computer space. A distance cutoff of 11 Å was used for the van der Waals and electrostatic interactions.

Implicit water solvation was simulated with Effective Energy Function 1 (EEF1) (Lazaridis and Karplus, 1999Go) implemented in the c27b1 version of CHARMM (Brooks et al., 1983Go). This implicit solvent model uses the CHARMM 19 polar hydrogen energy function (Neria et al., 1996Go) and evaluates the solvation energy of the protein based on the solvent exclusion volume of the protein atoms and a distance-dielectric constant. The ability of this model to attain a global balance between the intramolecular protein–protein interactions and the protein–solvent interactions in protein simulations was demonstrated by its ability to reproduce r.m.s.d.s similar to those in explicit solvent simulations for stable native protein conformations (Lazaridis and Karplus, 1999Go). Recent applications in protein folding/unfolding studies have further yielded results comparable to those of explicit solvent simulations (Lazaridis and Karplus, 1997Go; Inuzuka and Lazaridis, 2000Go). Interestingly, the electrostatic energy obtained in the EEF1 model correlates excellently with the Screened Coulumb Potential–Implicit Solvent model and reasonably well with PB energies (Hassan and Mehler, 2002Go). A time step of 2 fs was used throughout the EEF1 molecular dynamics simulations. Five multiple runs with different minimization steps (300–800) and different starting random seeds were carried out for each group of simulations.

The implicit solvation model, such as EEF1 used in this study, parameterizes static solvation effects (e.g. electrostatic and hydrophobic interactions). To consider further the dynamic effects of the missing water molecules on the proteins, we also simulated the unfolding of ß2-m with Langevin dynamics. The dynamic effect is incorporated in Langevin dynamics by a random collisions term (which models the thermal effect on the interaction of the protein with the solvent) and a friction term (which models the solvent viscosity effect). The Langevin dynamics simulations were carried out with the EEF1 force field combined with the multiple time-scales method (LN) (Barth and Schlick, 1998Go). Unlike the normal multiple time-scales schemes, LN merges the slow and fast time-scale forces via extrapolation rather than ‘impulses’ and allows larger time steps (Barth and Schlick, 1998Go). It has been shown that LN achieves very good agreement with small time step solutions of the Langevin equation in terms of thermodynamics, geometry and dynamics for proteins in vacuum and in large water systems (Barth and Schlick, 1998Go). The Langevin dynamics in LN were used here primarily to explore large-scale motion in a relatively low simulation temperature (400 K). The Langevin dynamics at 500 K produce similar results to those obtained from MD simulations at 500 K. The LN Langevin dynamics were run with the scheme of LN12 with a friction coefficient of 20 ps-1. The integrated time step in the LN12 setup includes 0.5 fs for the fast time step, 3.0 fs for the medium time step and 36 fs for the slow time step. Therefore, each dynamics step in the simulation reprensents a 36 fs time-scale. The forces from all bond stretching and angle bending motion are considered as fast time-scale motion. Forces from all dihedral and improper torsion motions are considered as medium time-scale motion.

As control and part of the examination, we simulated the native and {Delta}N6 ß2-ms at room temperature using the Langevin dynamics. In general, comparison of the backbone atoms of the simulated structures with the crystal structures yields an r.m.s.d. of ~2–2.5 Å. The inter-residue contacts are defined by backbone hydrogen bonding, with an H···O distance cutoff of 2.5 Å. Each contact map has all the hydrogen bonds in five simulation runs around a given time. In order to avoid a transient movement such as that which may be observed in a single snapshot, three frames (with t, t 1, t + 1) at given time t are combined. Thus, a single contact map contains hydrogen bonds from 15 snapshots. Backbone r.m.s.d.s are calculated using all backbone atoms with the crystal structure as the reference. Average r.m.s.d.s are calculated from the average of five r.m.s.d.s from five runs.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Thermal unfolding of native ß2-microglobulin with explicit water solvation

Three simulations are performed with explicit water solvation. First we simulate unfolding with the protein solvated with a 20 Å shell of water at 450 K. In the second and third simulations, the protein is solvated in 60x60x60 Å cubic box with periodic boundary conditions (PBC) at 500 and at 600 K, respectively. The total unfolding simulation time of ß2-m with the water shell solvation lasts for 1.9 ns.

The unfolding trajectory of the ß2-m indicates a hierarchical process (Figure 2A and B). In the first 0.5 ns, the protein three-dimensional structure is largely preserved, with only some local changes. Around 1.0 ns, the interactions between strands two (ß2) and six (ß6) are lost and the five-stranded ß-sheet (ß-strands ß1, ß2, ß4, ß5 and ß6) breaks into two parts: one with ß1 and ß2 and the other with ß4, ß5 and ß6 (Figure 2B). Since ß-strands ß2 and ß7 are linked with a disulfide bond, ß-strands ß1, ß2, ß3, ß7 and ß8 act as a domain (Figure 2B). As a result, we observe a clear ‘domain’ movement of the opening and flipping of the building block containing strands ß4, ß5 and ß6 from the rest of the protein (1.0–1.7 ns, Figure 2A and B).



View larger version (36K):
[in this window]
[in a new window]
 
Fig. 2. Backbone r.m.s.d. and snapshots from MD simulations with shell waters (A and B), PBC water at 500 K (C and D) and PBC water at 600 K (E and F).

 
The large domain movement was not observed in the PBC simulations. Unfolding processes under the PBC simulation protocol are characterized by a gradual melting of the local ß structure. As may be seen from the backbone r.m.s.d. trajectories in Figure 2, the common feature, especially in the earlier stage of unfolding [0–1.0 ns for shell water (Figure 2A) and PBC at 600 K (Figure 2E), 0–2.5 ns for PBC 500 K simulation (Figure 2C)], is the stability of the ß3–ß7 and ß2–ß6 interaction.

Figure 3 illustrates several important structural changes which were observed in this work. The domain motion depicted in Figure 3B appears in the water shell simulation. Figure 3C indicates a shift of ß4 to interact with ß3. Actually, the interaction of ß4 with ß6 in the crystal structure is very weak, with only a few interstrand contacts. Therefore, it is easy for ß4 to form a ß-sheet with ß3, as observed in the PBC 500 K simulation (Figure 2D). Figure 3D indicates the dissociation of ß7–ß8 and the subsequent shift of ß8 to interact with ß1, observed in the PBC 600 K simulation (Figure 2F). These movements appeared very often in the MD simulations with EEF1 force field (see next section, Table I).



View larger version (25K):
[in this window]
[in a new window]
 
Fig. 3. Important structural changes observed in the simulations. Note that some strands may be disrupted in the simulations. These are not reflected in these schemes. (A) The native structure with two ß-sheet layers: layer one with strands ß1, ß2, ß4, ß5 and ß6; layer two with strands ß3, ß7 and ß8. (B) Domain separation of strands ß4, ß5 and ß6 from the rest of protein. (C) Shifting of ß4 and ß5 to layer two. (D) Shifting of ß8 to layer one. (E) The changes in secondary structure from ß-strands to {alpha}-helices.

 
Thermal unfolding of native ß2-microglobulin with implicit water solvation

Twenty-two simulations were carried out for the native ß2-microglobulin with the implicit water solvation model (EEF1 force field). MD simulations with the CHARMM EEF1 force field were run at 450 and 500 K and Langevin dynamics with the CHARMM EEF1 force field were run at 400 and 500 K. With each simulation condition (MD/450 K, MD/500 K, LD/400 K and LD/500 K), five simulations are performed with different starting random seeds (Table I). Two simulations at room temperature are also carried out using the Langevin dynamics. The backbone r.m.s.d.s between the starting crystal structure and room temperature trajectories are around 2.0–2.5 Å.

Some selected snapshots from the unfolding simulations are shown in Figures 4 and 5. Figure 6 presents averaged r.m.s.d. trajectories for the simulations (averaged over five simulations). In Figures 7, 8 and 9, we analyze the changes in residue contacts during the simulations. We monitor the backbone hydrogen bonds as the index of residue contacts. Altogether, hydrogen bonds from five simulations at given times are projected on the contact map. For example, the native, 0.5 ns map in Figure 7 shows all the hydrogen bonds from snapshots taken at around 0.5 ns in the five molecular dynamics simulations at 450 K.



View larger version (35K):
[in this window]
[in a new window]
 
Fig. 4. Snapshots from 30 molecular dynamics simulations: (A) native ß2-m, 450 K at 4.0 ns; (B) truncated {Delta}N6 ß2-m, 450 K at 4.0 ns; (C) Lys-cut ß2-m, 450 K at 4.0 ns; (D) native ß2-m, 500 K at 0.5 ns; (E) {Delta}N6 ß2-m, 500 K at 0.5 ns; and (F) Lys-cut ß2-m, 500 K at 0.5 ns.

 


View larger version (39K):
[in this window]
[in a new window]
 
Fig. 5. Snapshots from 15 Langevin dynamics simulations with EEF1 force field for (A) native ß2-m at 400 K, 72 ns; (B) native ß2-m at 500 K, 3.6 ns; and (C) {Delta}N6 truncated ß2-m at 400 K, 72 ns.

 


View larger version (27K):
[in this window]
[in a new window]
 
Fig. 6. Average backbone r.m.s.d. from the trajectories of the simulations of native ß2-m [(A) MD simulations at 450 K; (B) MD simulations at 500 K; (C) Langevin dynamics simulations at 400 K; (D) Langevin dynamics simulations at 500 K], {Delta}N6 truncated ß2-m [(E) MD simulations at 450 K; (F) MD simulations at 500 K; (G) Langevin dynamics simulations at 400 K] and Lys-cut ß2-m [(H) MD simulations at 450 K; (I) MD simulations at 500 K]. Each r.m.s.d. line is averaged from five simulation runs.

 



View larger version (99K):
[in this window]
[in a new window]
 
Fig. 7. (A) and (B) contact (hydrogen bonds) maps from unfolding molecular dynamics simulation at 450 K for the native ß2-m (top), truncated {Delta}N6 ß2-m (middle row) and Lys-cut ß2-m (bottom).

 


View larger version (49K):
[in this window]
[in a new window]
 
Fig. 8. Contact (hydrogen bonds) maps from unfolding molecular dynamics simulation at 500 K for native ß2-m (top), truncated {Delta}N6 ß2-m (middle row) and Lys-cut ß2-m (bottom).

 


View larger version (50K):
[in this window]
[in a new window]
 
Fig. 9. Contact (hydrogen bonds) maps from unfolding Langevin dynamics simulation for native ß2-m at 400 K(top), truncated {Delta}N6 ß2-m at 400 K (middle row) and native ß2-m at 500 K (bottom).

 
The unfolding processes from each individual run show different dissociation and reformation behavior of the ß-strand contacts (for example, via the mechanisms shown in Figure 3C and D). This may be seen in the snapshots in Figures 4 and 5 and in the summary Table I. However, when we look at the five simulations combined, the consensus unfolding pattern appears. In the averaged r.m.s.d. trajectories for all native ß2-microglobulin simulations (Figure 6A–D), the ß2–ß6 interactions are the most stable, consistent with the PBC explicit water simulations. The ß3–ß7 interactions (which are also stable in the PBC simulations) are only stable in the molecular simulations at 450 K (Figure 6A).

The ß1–ß2 interactions, which are highly flexible in the PBC simulations, are relatively stable in the CHARMM EEF1 simulations. The underlying mechanism accounting for the ß1–ß2 stability is the dissociation of ß7–ß8 and the subsequent shift of ß8 to interact with ß1. The new ß1–ß8 interaction characterizes the structural changes from all four groups of simulations. This new ß-strand contact may be pictorially viewed in some snapshots in Figures 4 and 5. All contact maps (Figures 7, 8 and 9) show the appearance of such a new contact in the upper left corner.

As strand ß4 runs between the two layers, it may easily move between the layers. In the explicit water simulations, ß4 frequently extends to form a ß-sheet with strand ß3 in the second layer. The simulations also show a possible extension of the ß4–ß6 interactions (Table I).

Other important observed structural events are the changes of ß-strands to {alpha}-helices during the molecular dynamics simulations at 500 K (Table I). In the first run, the fragment of Ile91–Asp97 changed briefly to an {alpha}-helix at around 1.5 ns. In the second run, strands ß4 (Lys45–Ser51) and ß5 (Ser54–Ser60) partially changed to {alpha}-helices at around 0.42 ns. The Lys45–Gln51 fragment quickly changed back to a random coil; however, the Ser54–Trp60 {alpha}-structure region lasts a long time (one of the snapshots is shown in Figure 4D2, taken at 0.5 ns) to 1.0 ns. After 1.0 ns, only some of the hydrogen bonds exist within the Ser54–Trp60 fragment. However, at the end of the simulations at around 2.0 ns, the {alpha}-structure shifts down to the Asp58–Leu63 region. In the third run, ß3 (Pro33–Leu40) changes to an {alpha}-helix and lasts to 1.5 ns. In the fifth run, ß4 (Leu40–Ser48) changes to an {alpha}-helix at around 2.0 ns. The contact map in Figure 8 (native, 1.0 ns) shows the appearance of the {alpha}-helical contacts. Combined, these results clearly indicate that ß4 and ß5 easily change to {alpha}-helical structures.

Thermal unfolding of truncated {Delta}N6 ß2-microglobulin with implicit water solvation

Seventeen simulations were carried out for the {Delta}N6 ß2-microglobulin with the EEF1 force field. MD simulations were run at 450 and 500 K and Langevin dynamics were run at 400 K, all with multiple runs (five for each condition, Table I). Two room temperature simulations were also carried out using the Langevin dynamics to test the stability of truncated {Delta}N6 ß-m. However, we do not see a dramatic difference between the native and the truncated ß2-m at room temperature. The backbone r.m.s.d.s between the starting structure (assuming it is the same as the native crystal structure) and the room temperature trajectories are also around 2.0–2.5 Å. Room temperature simulations do not have enough energy to overcome the barrier around the starting conformation in our limited time simulation. However, high-temperature unfolding simulations do show the destabilization of the native structure due to the deletion.

The immediate consequence of the deletion of six residues from the N-terminus is the destabilization of ß1–ß2. As may be seen from the contact maps in Figures 7, 8 and 9, the ß1–ß2 contacts in the lower left corner disrupt and disappear more quickly than in the case of the native ß2-microglobulin. This is easily understandable.

The destabilization of ß1–ß2 has snowball effects on the stability of other related interactions. First, the ß2–ß6 association becomes much more flexible. The average r.m.s.d. over the trajectory for the ß2–ß6 is larger than that of the native ß2-m (Figures 6 and 10), especially in the Langevin dynamics simulations (Figure 6G). The contact maps also indicate that the ß2–ß6 interactions disrupt and disappear more quickly (Figures 7, 8 and 9), in particular the contact between the end of strand ß2 and the starting region of ß6.



View larger version (39K):
[in this window]
[in a new window]
 
Fig. 10. Comparison of average backbone r.m.s.d. of ß-strand separation in the MD simulations at 450 K. Each line is averaged over five simulations.

 
The ß3–ß7 interactions also appear to be weaker in the molecular dynamics simulations. However, they remain as the most stable interactions in the Langevin dynamics simulations (Figures 69) at 400 K.

An interesting feature is the stabilization of the ß7–ß8 interactions due to the N-terminal deletion. In the native ß2-m, the ß7–ß8 dissociates and subsequently ß8 shifts to interact with ß1. However, ß1 is very flexible in the {Delta}N6 truncated protein and is unable to disturb the ß7–ß8 interactions. Therefore, compared with the native ß2-m, the contact maps of the {Delta}N6 truncated ß2-m are also characterized by the lack of new ß1–ß8 contacts in the upper left corner and by the stabilization of the ß7–ß8 contacts.

The {Delta}N6 deletion also facilitates the conversion of ß-strands to {alpha}-helices. For the native ß2-m, an {alpha}-helix appears only in the 500 K simulations. However, for the {Delta}N6 truncated ß2-m, the {alpha}-helix appears already in the 450 K simulations (Table I). In the first 450 K simulation run, ß3 (Ile29–Glu38) changes to an {alpha}-helix at around 3.3 ns and this {alpha}-helix is stable to the end of this simulation run (Figure 4B1). In the second run, ß5 (Asp52–Ser59) changes to an {alpha}-helix at around 3.4 ns and it also lasts to the end of the simulations (Figure 4B2). In the third run, ß4 and ß5 (Asp46–Ser56) change to an {alpha}-helix at around 0.75 ns. The helix is stable to around 3.5 ns. ß3 (Pro27–Asp32) also changes to an {alpha}-helix at around 3.0 ns. In one of the 400 K Langevin dynamics simulations, ß5 (Leu47–Trp53) briefly adopts an {alpha}-helical conformation.

To summarize, the {Delta}N6 deletion destabilizes the native conformation and facilitates the ß-strand to {alpha}-helix transition.

Thermal unfolding of Lys-cut ß2-microglobulin with implicit water solvation and comparisons between the simulations

We simulated a variant of ß2-m with the cleavage between Lys57 and Asp58. Five molecular dynamics simulations were performed at each temperature (400 and 500 K). Surprisingly, except for the high flexibility of strands ß4 and ß5, the unfolding behavior of the Lys-cut ß2-m is very similar to the native ß2-m (Figures 4 and 610, Table I). This behavior differs dramatically from the {Delta}N6 deletion discussed above.

The position of the Lys57–Asp58 cut is in the ß-hairpin loop linking strands ß5 and ß6 (Figure 1C). In our earlier molecular dynamics simulations of a ß-hairpin from protein G, we showed that a covalently connected (uncut) ß-hairpin loop is crucial for ß-hairpin stability (Ma and Nussinov, 2000Go). Therefore, the high flexibility of strands ß4 and ß5 is within expectation due to the cut. However, it is unexpected that strands ß2 and ß6 remain stable.


    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Our simulations reproduced many of the experimentally observed structural changes when the native ß2-m is perturbed. The most striking agreement is in the ß-strand to {alpha}-helix transition. Upon treatment of wild-type ß2-m with acetonitrile or trifluoroethanol, the predominant ß2-structure of the native protein is partially lost and helical structures are obtained (Heegaard et al., 2001Go). In our simulations, several fragments (ß3, Ile29–Glu38; ß4, Leu40–Ser48; and ß5, Asp52–Ser59) consistently change to a helical structure under different simulation conditions. ß8 (Ile91–Asp97) changes to an {alpha}-helix briefly. A recent NMR study of ß2-m indicated a frequent sampling of helical conformations over the whole sequence in the acid-denatured state (Katou et al., 2002Go). A secondary structure query using the PHD server (Rost and Sander, 1993Go) for the sequence G29FHPPQIEIDLLKNGEKIKSEQSDL SFSKDWS60 only shows that I37DLL40 may be a candidate for a helical structure while the remainder of the sequence is favored for a loop structure. However, the prediction is insignificant given the brevity of that sequence. Identification of the region responsible for the ß-strand to {alpha}-helix transition aids in understanding amyloid formation of the ß2-m.

Many amyloidogenic peptides or proteins show ß-strand to {alpha}-helix or {alpha}-helix to ß-strand transitions prior to amyloid formation (Pan et al., 1993Go; Chiti et al., 1999Go; Gasset et al., 1992Go; Walsh et al., 1999Go; Takahashi et al., 2000Go; Ciani et al., 2002Go). Polyalanine, a typical sequence with a high {alpha}-helix propensity, also forms a ß-aggregation, as indicated by both experiments (Blondelle et al., 1996Go) and molecular dynamics simulations (Ma and Nussinov, 2002Go). Through a study of TFE-induced non-native {alpha}-helical structure in two ß-sheet-predominant proteins, ß-lactoglobulin and {alpha}-chymotrypsin, Dong et al. (Dong et al., 1998Go) found that the TFE-induced non-native {alpha}-helix structure in predominantly ß-sheet proteins is unstable and readily converts to an intermolecular ß-sheet aggregate. Others have also suggested that a defined {alpha}-helix structure in partially structured intermediates may favor a transition to ß-sheets and amyloid fibril formation (Chiti et al., 1999Go; Kayed et al., 1999Go; Rochet and Lansbury, 2000Go; Heegaard et al., 2001Go). Therefore, based on the ß-strand to {alpha}-helix transition, we may tentatively identify the region responsible for the intermolecular interaction leading to the polymerization as ß3, ß4 and ß5.

The conformational changes involved in amyloid formation are too complicated to understand from limited experimental and simulation results. Nevertheless, the current simulations hopefully provide some insight into the amyloid formation process. Native ß2-m forms amyloids, {Delta}N6 ß2-m increases amyloid formation and Lys-cut ß2-m does not form amyloids. Through comparisons of the conformational behavior of the native, {Delta}N6 and the Lys-cut ß2-m, using the three simulation methods, we attempt to deduce the conformational changes which are responsible for amyloid formation. Comparisons of the contact maps in Figures 79 and of the backbone r.m.s.d. trajectories in Figures 6 and 10 reveal increasing flexibility in the interactions of ß2–ß6 and ß3–ß7 in {Delta}N6 ß2-m.

NMR analysis shows that {Delta}N6 ß2-m is destabilized by 2.5 kcal/mol as compared with the intact protein. For the {Delta}N6 ß2-m, the loss of protection occurs in ß1, ß3 and part of ß2 (Esposito et al., 2000Go), in agreement with the simulations. Therefore, exposure of the previously buried strands ß2, ß3, ß6 and ß7 may also be responsible for the increasing amyloid formation in {Delta}N6 ß2-m.

NMR data (Esposito et al., 2000Go) also indicate that for {Delta}N6 ß2-m a similar ß-structure conformation is retained for ß6 and for the first segment of ß2 as compared with the native ß2-m. Since the last segment of ß2 is disturbed, both experiment and the present simulations suggest that parts of the ß2–ß6 interactions have been lost. Esposito et al. (Esposito et al., (2000Go) speculated that strand ß6 could be supported by pairing to strand ß5. This speculation receives strong support from our simulations. In the shell water simulation we observed a large domain movement (Figure 2B), in which ß5–ß6 act as one domain and separate from the rest of the protein. Experimentally, a partially structured species of ß2-m was found significantly populated under physiological conditions and involved in fibrilogenesis (Chiti et al., 2001Go). However, we do not have evidence to relate the specific unfolded conformation to experimental observation. Many occurrences of separation movements, on a smaller scale and involving only ß5 and part of ß6 pairing with ß5, appear in the simulations of {Delta}N6 ß2-m (for example, Figure 4B1). Based on the NMR analysis, Esposito et al. also suggested that strand ß5 is involved in intermolecular interactions that stabilize the association of {Delta}N6 ß2-m (Esposito et al., 2000Go).

No single experiment provides a definitive answer to the mechanism of amyloid formation for ß2-m. Limited proteolysis shows that the fibril core is comprised of residues 20–87 with strands (ß1 and ß8) unconstrained in the fibrillar polymer (Monti et al., 2002Go). A docking study of the conserved core (without strands ß1 and ß8) nicely produced the helical fibril structure for ß2-m (Benyamini et al., 2003Go). Stoppini et al. (Stoppini et al., 1997Go) found that amyloid formation may be inhibited by binding anti-(ß2-m) mAb to residues 92–99 (strand ß8) of ß2-m, whereas the binding to sequences 20–41 (strands ß2 and ß3) and 63–75 (strand ß6) have no effect on ß2-m fibrilogenesis. It appears that one may exclude strands ß2, ß3 and ß6 from participating in an intermolecular contact. Kozhukh et al. (Kozhukh et al., 2002Go) investigated peptides from ß2-m that form amyloids. They found that the isolated peptide Ser20–Lys41 (ß2–ß3) spontaneously formed amyloid fibrils. At the same time, they also found that the CD signal and the binding of the peptide Ser20–Lys41 and ß2-m are different. Therefore, regions other than Ser20–Lys41 in the intact ß2-m may also be involved in the ß2-sheet conformation in the fibrils. The identification of isolated ß2–ß3 to be amyloidogenic also agrees with the {alpha}-helix formation of ß3 in our simulation.

Two recent elegant NMR studies have revealed additional details regarding unfolding and amyloid formation of ß2-m. McParland et al. (McParland et al., 2002Go) measured the equilibrium denaturation of ß2-m at low pH. They found that the amyloid precursor is a non-cooperatively stabilized ensemble that retains most of the ß-strands. Hoshino et al. (Hoshino et al., 2002Go) compared the H–D exchange of amide protons of native ß2-m and its amyloid. Both studies point to the N- and C-terminal ß-strands as flexible and unprotected in the amyloid fibrils. Interestingly, strands ß4 and ß5, which are flexible in the native state, become highly protected in the amyloid fibril. On the other hand, whereas strands ß2 and ß7 are highly protected in the native state, in the amyloid the extent of their protection is unclear.

Combining the above experimental findings and simulations, the candidates responsible for intermolecular interactions of ß2-m could most likely be ß3, ß4 and ß5. The involvement of ß4 and ß5 is strongly supported both by the NMR analysis and by the current simulations (i.e. the {alpha}-helix transition and the movement of the paired ß5–ß6). It also explains why the Lys-cut ß2-microglobulin does not form amyloids. As indicated in our simulations of Lys-cut ß2-m, ß5 is the most flexible region, making a ß5 pairing with ß6 impossible after cutting at Lys57–Asp58. The involvement of ß3 is supported by its high H–D exchange protection in the amyloid fibril (Hoshino et al., 2002Go), by amyloid formation of the short peptide fragment (Kozhukh et al., 2002Go) and by our observation of its {alpha}-helix transition in the simulation. The reason that the antibody binding of sequences 20–41 (strands ß2 and ß3) does not prevent amyloid formation (Stoppini et al., 1997Go) may be due to the {alpha}-helix transition which escapes the antibody binding before amyloid formation.

The polymerization of proteins into amyloids with regular structures argues for association of monomers which contain some defined elements of structure and which can make intermolecular contacts (Sunde et al., 1997Go; Sunde and Black, 1998Go). It was suggested that amyloid formation can be explained remarkably well through a ‘domain swapping’ event, where the swapped segment is either a ß-hairpin or a conformation which can assume a ß-hairpin structure (Sinha et al., 2001Go). In this way, most of the ß-structure present in the native protein may be preserved and extended in the resulting amyloids. To permit such a mechanism, there should exist a structural motif in the native protein, facilitating structural plasticity. For example, it was postulated that the ‘loss’ of ß-strand from the transthyretin structure could uncover an amyloidogenic determinant which allows its ß-sheet structure to make intermolecular ß-type hydrogen bonds and thereby propagate indefinite sheets of the kind proposed to exist in the fibrils (Kelly and Lansbury, 1994Go; Serpell and Blake, 1994Go). The mechanism that we propose here follows a similar pathway.

Conclusions

In this study, we examined the unfolding processes of ß2-microglobulin and two related variants (one with N-terminal hexapeptide deletion {Delta}N6 and the other with Lys57–Asp58 cleavage) by high-temperature molecular dynamics simulations. When we perturb the native conformation of ß2-m, our simulations reproduce many of the experimentally observed structural changes. The clearest agreement is in the ß-strand to {alpha}-helix transition. In our simulations, strands ß3, ß4 and ß5 consistently change to {alpha}-helices and ß8 changes briefly to an {alpha}-helix. The results of our simulations provide both a mechanism and atomic detail for the experimentally observed ß-strand to {alpha}-helix change for ß2-m and strongly support the proposition that ß–{alpha}–ß transitions may be an important step in amyloid formation.

By comparing the conformational behavior of the native, {Delta}N6 and the Lys-cut ß2-m, using three simulation methods, we identified the consensus conformational changes that differentiate between the native ß2-m and its two variants. The {Delta}N6 deletion destabilizes the native conformation and facilitates the ß-strand to {alpha}-helix transition. The increasing flexibility of ß2–ß6 and ß3–ß7 which we observe agrees with the NMR data. The main effect of the removal of the N-terminal hexapeptide is the increasing separation between strands ß2 and ß6, while the lysine cleavage only increases the flexibility of strand ß5 and has no effect on the interactions between strands ß2 and ß6. Since the native ß2-m forms amyloids, {Delta}N6 ß2-m increases amyloid formation and the Lys-cut ß2-m does not form amyloids, the simultaneous dislocation of strands ß5 and ß6 may be responsible for the polymerization. However, it is difficult to relate conformational change directly to amyloid formation based on current experimental knowledge and simulated results with a limited time-scale.


    Acknowledgements
 
We thank Dr Jacob V.Maizel for encouragement and for helpful discussions. We thank Dr Neeti Sinha for helpful discussions. The computation times were provided by the National Cancer Institute’s Frederick Advanced Biomedical Supercomputing Center and utilized the high-performance computational capabilities of the Biowulf/LoBoS3 cluster at the National Institutes of Health, Bethesda, MD. The research of R.Nussinov in Israel has been supported in part by a Ministry of Science grant, by a Magnet grant and by the Center of Excellence in Geometric Computing and its Applications funded by the Israel Science Foundation (administered by the Israel Academy of Sciences). This project has been funded in whole or in part with Federal funds from the National Cancer Institute, National Institutes of Health, under contract number NO1-CO-12400. The content of this publication does not necessarily reflect the view or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products or organizations imply endorsement by the US Government.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Barth,E. and Schlick,T. (1998) J. Chem. Phys., 109, 1617–1632.[CrossRef][ISI]

Becker,J.W. and Reeke,G.N.,Jr. (1985) Proc. Natl Acad. Sci. USA, 82, 4225–4229.[Abstract]

Bellotti,V., Stoppini,M., Mangione,P., Sunde,M., Robinson,C., Asti,L., Brancaccio,D. and Ferri,G. (1998) Eur. J. Biochem., 258, 61–67.[Abstract]

Bellotti,V., Gallieni,M., Giorgetti,S. and Brancaccio,D. (2001) Semin. Dial., 14, 117–122.[CrossRef][ISI][Medline]

Benyamini,H., Gunasekaran,K., Wolfson,H. and Nussinov,R. (2003) J. Mol. Biol., 330, 159–174.[CrossRef][ISI][Medline]

Bernstein,F.C., Koetzle,T.F., Williams,G.J.B., Meyer,E.F., Brice,M.D., Rodgers,J.R., Kennard,O., Shimanouchi,T. and Tasumi M. (1977) J. Mol. Biol., 112, 535–542.[ISI][Medline]

Blondelle,S.B., Forood,B., Houghten,R.A. and Pérez-Payá,E. (1996) Biochemistry, 36, 8393–8400.[CrossRef][ISI]

Brooks,B.R., Bruccoleri,R.E., Olafson,B.D., Sate,D.J., Swaminathan,S. and Karplus,M. (1983) J. Comput. Chem., 4, 187–217.[ISI]

Carrell,R.W. and Lomas,D.A. (1997) Lancet, 350, 134–138.[CrossRef][ISI][Medline]

Chiti,F., Webster,P., Taddei,N., Clark,A., Stefani,M., Ramponi,G. and Dobson,C.M. (1999) Proc. Natl Acad. Sci. USA, 96, 3590–3594.[Abstract/Free Full Text]

Chiti,F., Lorenzi,E.D., Grossi,S., Mangione,P., Giorgetti,S., Caccialanza,G., Dobson,C.M., Merlini,G., Ramponi,G. and Bellotti,V. (2001) J. Biol. Chem., 276, 46714–46721.[Abstract/Free Full Text]

Ciani,B., Hutchinson,E.G., Sessions,R.B. and Woolfson,D.N. (2002) J. Biol. Chem., 227, 10150–10155.[CrossRef]

Dong,A., Matsuura,J., Manning,M.C. and Carpenter,J.F. (1998) Arch. Biochem. Biophys., 355, 275–281.[CrossRef][ISI][Medline]

Esposito,G. et al. (2000) Protein Sci., 9, 831–845.[Abstract]

Floege,J. and Ketteler,M. (2001) Kidney Int., 59, S-164–S-171.[ISI]

Gasset,M., Baldwin,M.A., Lloyd,D.H., Gabriel,J.M., Holtzman,D.M., Cohen,F., Fletterick,R. and Prusiner,S.B. (1992) Proc. Natl Acad. Sci. USA, 89, 10940–10944.[Abstract]

Gilquin,B., Guilbert,C. and Perahia,D. (2000) Proteins, 41, 58–74.[CrossRef][ISI][Medline]

Harrison,P.M., Bamborough,P., Daggett,V., Prusiner,S. and Cohen,F.E. (1997) Curr. Opin. Struct. Biol., 7, 53–59.[CrossRef][ISI][Medline]

Hassan,S.A. and Mehler,E.L. (2002) Proteins, 47, 45–61.[CrossRef][ISI][Medline]

Heegaard,N.H., Sen,J.W., Kaarsholm,N.C. and Nissen,M.H. (2001) J. Biol. Chem., 276, 32657–32662.[Abstract/Free Full Text]

Hoshino,M., Katou,H., Hagihara,Y., Hasegawa,K., Naiki,H. and Goto,Y. (2002) Nat. Struct. Biol., 9, 332–336.[CrossRef][ISI][Medline]

Inuzuka,Y. and Lazaridis,T. (2000) Proteins, 41, 21–32.[CrossRef][ISI][Medline]

Katou,H., Kanno,T., Hoshino,M., Hagihara,Y., Tanaka,H., Kawai,T., Hasegawa,K., Naiki,H. and Goto,Y. (2002) Protein Sci., 11, 2218–2229.[Abstract/Free Full Text]

Kayed,R., Bernhagen,J., Greenfield,N., Sweimeh,K., Brunner,H., Voelter,W. and Kapurniotu,A. (1999) J. Mol. Biol., 287, 781–796.[CrossRef][ISI][Medline]

Kelly,J.W. and Lansbury,P.T. (1994) Amyloid, 1, 186–205.

Kozhukh,G.V., Hagihara,Y., Kawakami,T., Hasegawa,K., Naiki,H. and Goto,Y. (2002) J. Biol. Chem., 277, 1310–1315.[Abstract/Free Full Text]

Lazaridis,T. and Karplus,M. (1997) Science, 278, 1928–1931.[Abstract/Free Full Text]

Lazaridis,T. and Karplus,M. (1999) Proteins, 35, 133–152.[CrossRef][ISI][Medline]

Ma,B. and Nussinov,R. (2000) J. Mol. Biol., 296, 1091–1104.[CrossRef][ISI][Medline]

Ma,B. and Nussinov,R. (2002) Protein Sci., 11, 2335–2350.[Abstract/Free Full Text]

Ma,B., Wolfson,H. and Nussinov,R. (2001) Curr. Opin. Struct. Biol., 11, 364–369.[CrossRef][ISI][Medline]

Maple,J.R., Hwang,M.J., Jalkanen,K.J., Stockfisch,T.P. and Hagler,A.T. (1998) J. Comput. Chem., 19, 430–458.[CrossRef][ISI]

McParland,V.J., Kalverda,A.P., Homans,S.W. and Radford,S.E. (2002) Nat. Struct. Biol., 9, 326–331.[CrossRef][ISI][Medline]

Monti,M., Principe,S., Giorgetti,S., Mangione,P., Merlini,G., Clark,A., Bellotti,V., Amoresano,A. and Pucci,P. (2002) Protein Sci., 11, 2362–2369.[Abstract/Free Full Text]

Neria,E., Fischer,S. and Karplus,M. (1996) J. Chem. Phys., 105, 1902–1921.[CrossRef][ISI]

Pan,K.M. et al. (1993) Proc. Natl Acad. Sci. USA, 90, 10962–10966.[Abstract]

Rochet,J.C. and Lansbury,P.T.,Jr. (2000) Curr. Opin. Struct. Biol., 10, 60–68.[CrossRef][ISI][Medline]

Rost,B. and Sander,C. (1993) J. Mol. Biol., 232, 584–599.[CrossRef][ISI][Medline]

Serpell,L.C. and Blake,C.C.F. (1994) In Kisilevsky,R., Benson,M.D., Frangione,B., Gauldie,J., Muckle,T.J. and Youngs,I.D. (eds), Amyloid and Amyloidosis. Parthenon, New York, pp. 447–450.

Sinha,N., Tsai,C.J. and Nussinov,R. (2001) Protein Eng., 14, 93–103.[Abstract/Free Full Text]

Smith,D.P. and Radford,S.E. (2001) Protein Sci., 10, 1775–1784.[Abstract/Free Full Text]

Stoppini,M., Bellotti,V., Mangione,P., Merlini,G. and Feri,G. (1997) Eur. J. Biochem., 249, 21–26.[Abstract]

Sunde,M. and Blake,C.C.F. (1998) Q. Rev. Biophys., 31, 1–39.[CrossRef][ISI][Medline]

Sunde,M., Serpell,L.C., Bartlam,M., Pepys,M.B., Fraser,P.E. and Blake,C.C.F. (1997) J. Mol. Biol., 273, 729–739.[CrossRef][ISI][Medline]

Takahashi,Y., Ueno,A. and Mihara,H. (2000) Struct. Fold. Des., 8, 915–925.[ISI][Medline]

Walsh,D.M., Hartley,D.M., Kusumoto,Y., Fezoui,Y., Condron,M.M., Lomakin,A., Benedek,G.B., Selkoe,D.J. and Teplow,D.B. (1999) J. Biol. Chem., 274, 25945–25952.[Abstract/Free Full Text]

Received September 18, 2002; revised June 23, 2003; accepted June 25, 2003.





This Article
Abstract
FREE Full Text (PDF)
Alert me when this article is cited
Alert me if a correction is posted
Services
Email this article to a friend
Similar articles in this journal
Similar articles in ISI Web of Science
Similar articles in PubMed
Alert me to new issues of the journal
Add to My Personal Archive
Download to citation manager
Search for citing articles in:
ISI Web of Science (4)
Request Permissions
Google Scholar
Articles by Ma, B.
Articles by Nussinov, R.
PubMed
PubMed Citation
Articles by Ma, B.
Articles by Nussinov, R.