Human Islet Amyloid Polypeptide N-Terminus Fragment Self-Assembly: Effect of Conserved Disulfide Bond on Aggregation Propensity
Amyloid formation by human islet amyloid polypeptide (hIAPP) has long been implicated in the pathogeny of type 2 diabetes mellitus (T2DM) and failure of islet transplants, but the mechanism of IAPP self-assembly is still unclear. Numerous fragments of hIAPP are capable of self-association into oligomeric aggregates, both amyloid and non-amyloid in structure. The N-terminal region of IAPP contains a conserved disulfide bond between cysteines at position 2 and 7, which is important to hIAPP’s in vivo function and may play a role in in vitro aggregation. The importance of the disulfide bond in this region was probed using a combination of ion mobility-based mass spectrometry experiments, molecular dynamics simulations, and high-resolution atomic force microscopy imaging on the wildtype 1-8 hIAPP fragment, a reduced fragment with no disulfide bond, and a fragment with both cysteines at positions 2 and 7 mutated to serine. The results indicate the wildtype fragment aggregates by a different pathway than either comparison peptide and that the intact disulfide bond may be protective against aggregation due to a reduction of inter-peptide hydrogen bonding.
KeywordsAmyloid IAPP Islet amyloid polypeptide Diabetes T2DM Type 2 diabetes mellitus human Islet Amyloid Polypeptide hIAPP Ion mobility Electrospray ionization Peptide Atomic force microscopy Replica exchange molecular dynamics
Increasing evidence correlates the process of amyloid formation by human islet amyloid polypeptide (hIAPP) with pathogenic β-cell apoptosis in diabetes mellitus type II (T2DM) [1, 2, 3, 4, 5, 6, 7, 8]. hIAPP is a 37-residue peptide co-secreted with insulin from pancreatic β-cells, which normally plays an adaptive role in glucose metabolism. In T2DM this peptide is overexpressed and misfolds from its natively disordered state [8, 9, 10, 11], forming pervasive amyloid plaques in the pancreas. Amyloid formation by hIAPP also contributes to the failure of islet transplants [12, 13]. As with other amyloid diseases, hIAPP plaques are composed of macroscopic, structured fibrils containing stacked parallel protein β-sheets [1, 14]. Recent findings suggest that it is not the plaques but early soluble oligomers that are the most pathogenic agent [2, 3, 15, 16]. It is therefore critical to understand early oligomer assembly and structure, a process hampered by the transient nature of the soluble oligomers. Ion mobility spectrometry-mass spectrometry (IMS-MS) has proven to be a powerful tool for investigating the distribution of these low-order amyloid oligomers [17, 18, 19, 20, 21]. Previous IMS-MS studies have revealed key differences between the oligomers formed by hIAPP and the oligomers formed by nontoxic rat IAPP [22, 23]. IMS-MS has also been used to study the inhibition of hIAPP amyloid formation in vitro by insulin and small molecules [20, 21, 24].
The aggregation dynamics of hIAPP peptide segments have been studied for their potential as aggregation nucleation sites while modified fragments serve as promising inhibitors of hIAPP aggregation [25, 26, 27, 28, 29, 30]. Much attention has focused on hIAPP20-29 (consisting of the 20–29 residue region, SNNFGAILSS) and hIAPP22-29 as early work linked the primary sequence in this region to hIAPP amyloidogenicity. Subsequent work showed the situation is more complex but the 22-29 region is still considered to be a key contributor to hIAPP aggregation [31, 32, 33]. Several other IAPP segments have been found capable of self-association, including the N-terminal region [27, 34, 35]. This region, which contains an intramolecular disulfide bond between cysteine residues 2 and 7, is thought to be important to hIAPP’s in vivo activity [36, 37, 38]. Additionally, while hIAPP has been found to endogenously decrease insulin secretion from β-cells, the introduction of hIAPP8-37 had an antagonistic effect on hIAPP-mediated insulin inhibition . There are conflicting reports on the role of the N-terminus in hIAPP aggregation; early studies showed that amyloid formation by an 8-37 fragment of hIAPP occurs more rapidly in buffer but more slowly in the presence of 1.5% hexafluoroisopropanol (HFIP) [40, 41]. In either case, the effect was modest but scanning transmission electron microscopy (STEM) appears to indicate that truncation changes the morphology of the fibrils . A mutant form of hIAPP, with alanine substitutions at both cysteine residues, positions 2 and 7, showed a modest increase in the nucleation lag-phase compared with wildtype hIAPP . A study by Cope et al. has shown that the hIAPP1-8 peptide fragment is capable of non-amyloid aggregation into large macroscopic structures and that cleavage of the disulfide bond apparently inhibits this aggregation process . Given both the physiological and biophysical significance of the disulfide bond in hIAPP, understanding the role of the N-terminal region is important to the overall picture of IAPP aggregation and pathology.
Materials and Methods
Peptide Synthesis and Preparation
All peptides were synthesized using Fmoc-based microwave-assisted methods and purified by reverse-phase HPLC as previously described [50, 51]. All peptides contain an amidated C-terminus. Samples were dissolved in 100% hexafluoroisopropanol (HFIP) (Sigma-Aldrich, St. Louis, MO, USA) to a final stock concentration of 1 mM. Aliquots of the stock were lyophilized and redissolved in 1:1 methanol:water for final peptide concentrations of 50–100 μM. In order to reduce the disulfide bond between cysteine residues 2 and 7, tris(2-carboxyethyl)phosphine (TCEP) (Alfa Aesar, Ward Hill, MA, USA) was added at a 10:1 excess to peptide solutions and briefly incubated before measurements were taken. Reduction of the disulfide bond was verified via quadrupole time-of-flight mass spectrometry (Supplementary Figure S4). All solvents had their pH adjusted to physiological pH (7.4) using concentrated ammonium hydroxide and acetic acid unless otherwise noted.
Ion Mobility Experiments
An in-depth description of the IMS-MS instrumentation has been given previously . In summary, samples were loaded into gold-coated borosilicate glass capillaries that had been pulled to a fine point using an in-house capillary puller (Sutter Instrument Co., Novato, CA, USA). Using a nanoelectrospray ionization source (nano-ESI), ions are pulled from the capillary under the influence of a voltage differential and channeled through a small (~70 μm) orifice into an ion funnel. The funnel then collects and focuses the ions; these can either be continuously funneled (for mass spectra) or pulsed (for ion mobility) into a 4.5-cm drift cell filled with ~3.5 Torr of helium gas. Under the influence of a weak electric field, ions are drifted through the cell, mass analyzed by a quadrupole, and detected by an electron multiplier detector. Total time for analysis is between 400 and 2000 μs.
Explicit solvent temperature-REMD (T-REMD) simulations were performed for the wildtype 1-8 residue fragment (WT1-8 ox, containing a disulfide bond between the second and seventh cysteine residues), the reduced 1-8 residue fragment (WT1-8 red), and the cysteine → serine C2S/C7S mutant fragment (C2S/C7S1-8) using the GROMACS 4.5.3 package and the all-atom Optimized Potentials for Liquid Simulations (OPLS-AA) force field in TIP3P water [54, 55, 56]. The peptide monomers have charged termini throughout the simulation, assuming neutral pH. The initial structure was minimized using the steepest decent method and solvated in a TIP3P water box. The size of the solvated cubic system was 4.50 nm for peptide fragment dimer and tetramer simulations. In each system, negative charge chlorine ions were added to neutralize the system. Solvent and volume equilibration simulations in NPT ensemble (T = 300 K and P = 1 bar) were performed to optimize the box size, followed by 6-ns NVT equilibration at 300 K. The LINCS algorithm was employed to constrain bonds between heavy atoms and hydrogen , and the SETTLE algorithm was used for water molecules . These constraints allowed an integration time step of 2.0 fs. The electrostatic and dispersion forces were computed with a real space cutoff of 1.2 nm, and particle mesh Ewald method was used to treat long-range electrostatics . The temperature was maintained by the Nose-Hoover thermostat . The temperature and pressure coupling constants were 0.1 and 1.0 ps, respectively. The equations of motion were integrated according to the leap-frog algorithm.
The initial guess for temperatures in T-REMD simulations (24 replicas for dimer and 32 replicas for tetramers) was taken from Patriksson and Spoel’s temperature predictor (http://folding.bmc.uu.se/remd/index.php) and then adjusted to obtain the exchange rate of approximately 20%–25% . Temperature lists can be found in the supporting information.
Each replica was equilibrated at the desired temperature for 12 ns before the production run for T-REMD began. Exchanges between replicas were attempted every 2 ps. The production run was 300-ns long per replica for dimers and 400-ns per replica for tetramers. Only the last 150-ns data of the trajectory at 299 K were subjected to analysis (Supplementary Figure S5). Clustering was performed using the Daura algorithm . In order to obtain theoretical cross sections, 50 structures per cluster were subjected to a short minimization in the gas phase to mimic the desolvation process after the electrospray ionization process. After that, the theoretical cross sections were computed using the projected superposition approximation (PSA) method available at (http://luschka.bic.ucsb.edu:8080/WebPSA/index.jsp) and the average values were reported [63, 64].
Atomic Force Microscopy
AFM experiments were carried out in tapping mode in air using an MFP-3D Atomic Force Microscope (Asylum Research, Goleta, CA, USA). A silicon probe was used with a cantilever spring constant of 7 N/m and a resonant frequency of 155 kHz (MikroMasch USA, Lady’s Island, SC, USA). Protein incubation conditions matched those used in the IMS-MS experiments. Oxidized and mutant peptide fragment samples were incubated in 1:1 methanol:water; chemically reduced peptide samples were incubated with TCEP reducing agent at a 10:1 ratio of reducing agent to peptide. Five μL aliquots were removed at defined times over the course of incubation and deposited on freshly cleaved V1-grade mica (TedPella, Redding, CA, USA). The samples were then dried in a vacuum desiccator. Height analysis of globular aggregates was performed using a mask at 1 nm to the image and adjusting to include all aggregates in the image. A particle analysis routine within the MFP-3D software was then applied.
Results and Discussion
Early Oligomer Formation Probed by IMS-MS Reveals a Distinct Pathway for the Aggregation of WT1-8ox Compared to Those of WT1-8red and C2S/C7S1-8
Collision Cross Section in Å2 of IAPP Fragments in the z/n = +1/1 and +1/2 ATDs*
Oxidized WT 1-8
C2S/C7S mutant 1-8
AFM Reveals WT1-8red and C2S/C7S1-8 Show Opposite Morphological Transformations
As seen in Figure 6b, c, both WT1-8 red and C2S/C7S1-8 rapidly form highly ordered aggregate structures (Figure 6, 5 min), consistent with both the higher order oligomerization seen in the IMS-MS experiments and the extensive intermolecular hydrogen bonding predicted by REMD. C2S/C7S1-8 fibrils of sizes ranging in width from 1 nm at the smallest to 10 nm at the largest were observed after just 5 min of incubation (Figure 6c). At longer incubation times, these fibrils appear to self-associate resulting in an interwoven morphology (Figure 6c, 30 min) that then rearranges into short, needle-like aggregates (Figure 6c, 60 min). WT1-8 red also formed ordered aggregates rapidly; however, its time evolution differs from C2S/C7S1-8. Short, closely associated fibril structures are favored at short incubation times and more extended fibrils favored at longer incubation times (Figure 6b). After 1 h, both WT1-8 red and C2S/C7S1-8 exhibited feature lengths significantly shorter than WT1-8 ox. This could explain the apparent lack of aggregation observed in prior studies for WT1-8 red, given that structures of this size would not be visible under a standard light microscope . Again it is intriguing that the peptides for which REMD predicted higher intermolecular bonding appear to follow a discrete aggregation pathway disparate from WT1-8 ox, a result consistent with IMS-MS data.
The presence of the disulfide bond in the N-terminal fragment does not completely inhibit self-aggregation of the fragment, but reduction of the disulfide bond or its removal by mutation increases the rate of aggregation. Overall, IMS-MS, REMD simulations, and AFM indicate WT1-8 ox is capable of forming stable aggregates along an entirely different pathway than both WT1-8 red and C2S/C7S1-8. WT1-8 red and C2S/C7S1-8 form extended oligomer conformations with high intermolecular hydrogen bonding (Figures 4 and 5). AFM images recorded at short incubation times corroborate these conclusions, with WT1-8 red and C2S/C7S1-8 showing extensive aggregation and rearrangement. Long-term macroscopic fibril formation varies drastically between the peptides, WT1-8 ox prefers extended fibril bundling seeded by globular aggregates; WT1-8 red forms short fibrils of varying widths, and C2S/C7S1-8 favors densely packed needle-like structures. These observations, coupled with the REMD results, suggest that the cleavage of the disulfide bond has an agonistic effect toward oligomerization of the full hIAPP peptide by increasing the probability of intermolecular hydrogen bonding and β-sheet formation. Perhaps the most important conclusion we make is that the 2-7 disulfide bond at the N-terminus limits the aggregation of the peptide relative to both the reduced and mutated peptides. This occurs primarily due to a significant reduction in intermolecular hydrogen bonds relative to the noncyclic peptides and suggests the disulfide bond may actually protect full-length hIAPP from amyloid formation. This central conclusion is at odds with the conclusion of Cope et al. , who concluded that “loop–loop” interaction might enhance hIAPP oligomerization and could be a target for therapeutic intervention.
The authors thank William Wonderly for his assistance with sample preparation. Support from the National Science Foundation under grant CHE-1301032 (M.T.B.) and the National Institutes of Health GM078114 (D.P.R.) is gratefully acknowledged. A.G.W. was supported in part by a GAANN fellowship from the Department of Education.
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