Make Illumina part of your DNA Sign up for PNAS Online eTocs
Link: Info for AuthorsLink: Editorial BoardLink: AboutLink: SubscribeLink: AdvertiseLink: ContactLink: Sitemap Link: PNAS Home
Proceedings of the National Academy of Sciences
Link: Current Issue "" Link: Archives "" Link: Online Submission ""  Link: Advanced Search

Published online on July 17, 2007, 10.1073/pnas.0703306104
PNAS | July 24, 2007 | vol. 104 | no. 30 | 12341-12346


This Article
Right arrow Abstract Freely available
Right arrow Figures Only
Right arrow Full Text (PDF)
Right arrow Supporting Information
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a colleague
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My File Cabinet
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via CrossRef
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ruschak, A. M.
Right arrow Articles by Miranker, A. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ruschak, A. M.
Right arrow Articles by Miranker, A. D.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg  
What's this?

 Previous Article  | Table of Contents |  Next Article 

BIOLOGICAL SCIENCES / BIOPHYSICS
Fiber-dependent amyloid formation as catalysis of an existing reaction pathway

Amy M. Ruschak* and Andrew D. Miranker{dagger},{ddagger}

Departments of *Chemistry and {dagger}Molecular Biophysics and Biochemistry, Yale University, 260 Whitney Avenue, New Haven, CT 06520-8114

Edited by Alan R. Fersht, University of Cambridge, Cambridge, United Kingdom, and approved June 5, 2007 (received for review April 10, 2007)


   Abstract
Top
 Abstract
Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 
A central component of a number of degenerative diseases is the deposition of protein as amyloid fibers. Self-assembly of amyloid occurs by a nucleation-dependent mechanism that gives rise to a characteristic sigmoidal reaction profile. The abruptness of this transition is a variable characteristic of different proteins with implications to both chemical mechanism and the aggressiveness of disease. Because nucleation is defined as the rate-limiting step, we have sought to determine the nature of this step for a model system derived from islet amyloid polypeptide. We show that nucleation occurs by two pathways: a fiber-independent (primary) pathway and a fiber-dependent (secondary) pathway. We first show that the balance between primary and secondary contributions can be manipulated by an external interface. Specifically, in the presence of this interface, the primary mechanism dominates, whereas in its absence, the secondary mechanism dominates. Intriguingly, we determine that both the reaction order and the enthalpy of activation of the two nucleation processes are identical. We interrogate this coincidence by global analysis using a simplified model generally applicable to protein polymerization. A physically reasonable set of parameters can be found to satisfy the coincidence. We conclude that primary and secondary nucleation need not represent different processes for amyloid formation. Rather, they are alternative manifestations of the same, surface-catalyzed nucleation event.

amylin | fibers | islet amyloid polypeptide | nucleation

The noncovalent, fibrous self-assembly of proteins, including hemoglobin S, actin, microtubules, and amyloid fibers, occurs by a nucleation-dependent mechanism (1–5). If polymer product formation is monitored as a function of time in a reaction where all of the protein is initially monodisperse, there is a lag phase in which little product is formed, followed by a period of rapid growth (2). Such observations are consistent with a rate-limiting step in which change occurs to a high-energy intermediate known as the nucleus. In many systems such as actin, collagen, and microtubules, the nucleus is modeled as an oligomeric species that is in a highly unfavorable equilibrium with monomer (1, 3–6). The nucleus may also be a high-energy conformation of monomer, such as that reported for polyglutamine (7). Regardless, the rate-limiting step involves a nucleus because it is defined as the species with the highest free energy and therefore lowest population (1, 4).

This model of nucleation alone cannot account for the high apparent cooperativity of conversion observed in many systems. Historically, hemoglobin S was the first biological polymerization reaction to demonstrate a transition time that is much shorter than its preceding lag phase (8). This same phenomenon is commonly reported for amyloid systems including islet amyloid polypeptide (IAPP) from type II diabetes (9), Abeta from Alzheimer's (10), PrP from the mammalian prion (11), and Sup35 from the yeast prion (12, 13), as well as model systems such as insulin (14). In such instances, a secondary (2° or fiber-dependent) mechanism of nucleation, in addition to the primary (1° or fiber-independent) one, is invoked to describe the kinetic profile. Indeed, polymer-dependent nucleation is fundamental not only to basic understanding of peptide physical chemistry but also to the capacity of these proteins to give rise to disease. For example, the relative infective capacity of mammalian (11) and yeast (13) prions and has been ascribed to intrinsic and environmental factors affecting 2° nucleation.

The nature of fiber-dependent nucleation is not understood for most biological polymers, particularly amyloid. There are three generally accepted possibilities for generating new ends from existing fibers. One is spontaneous or induced breakage (scission) of an existing polymer. This is the generally asserted mechanism for amyloid and is consistent with measures of fiber length (13) and tensile strength (15). Another possible mechanism is branching (1); however, the absence of imaging evidence for this makes it an unlikely mechanism for amyloid formation. Finally, it has been modeled as lateral nucleation from the walls of the polymers for hemoglobin S (8). In all cases, mechanisms constitute structurally and energetically distinct processes from 1° nucleation.

In this work, we determine the nature of nucleation processes for a peptide derived from the amyloidogenic protein IAPP. IAPP, also known as amylin, is a hormone cosecreted with insulin by the beta-cells of the pancreas. In type II diabetes, IAPP forms amyloid deposits that are correlated with beta-cell death (16). Recent transgenic models, e.g., the HIP rat, strongly support a role for IAPP in diabetic pathology (17). Residues 20–29 of IAPP, SNNFGAILSS, referred to here as IAPP20-29, have previously been shown to form amyloid independently of the rest of the sequence (18–20). Here, we study the kinetics of assembly of the cationic form of the peptide (amidated C terminus). Previous work has shown that full-length IAPP forms amyloid by both fiber-independent and fiber-dependent pathways (9). Here, we show that the 2° mechanism of nucleation also plays a prominent role in the fibrous assembly of IAPP20-29. We then determine the origins of this pathway and relate it to 1° nucleation.


   Results
Top
 Abstract
 Results
Discussion
 Materials and Methods
 Acknowledgements
 References
 
To define the processes governing the fibrous assembly of IAPP20-29, we first determine the minimum number of nucleation pathways necessary to describe the kinetics of assembly. We then establish solution conditions for which alternative pathways are dominant. Then, for each pathway, we characterize the reaction order and enthalpy of the transition state barrier of the rate-limiting step.

Nucleation of IAPP20-29 fibers occurs on surfaces. This is evident by comparing the kinetics of fiber formation reactions that differ only in the extent of filtration used to prepare reaction buffer. Fiber formation reactions are initiated by diluting a DMSO stock solution of IAPP20-29 (10 mM) into aqueous buffer. The buffer is prepared by using a filtered (0.2 µm) 10x stock diluted with water that is either nominally free of particulates (Milli-Q; Millipore, Billerica, MA) or has had an additional 0.2-µm filtration step applied immediately before use. Kinetics are then assessed by either 90° light scattering or the change in fluorescence of an introduced dye, thioflavin T (ThT) (21). Under the former conditions, the kinetic profile of a reaction with 700 µM IAPP20-29 is 50% complete (t50) in 3,400 ± 700 sec (Fig. 1A). In marked contrast, additional filtration only to the water results in a t50 of 32,000 ± 12,000 sec. This effect is not a consequence of the filter or the source of water [supporting information (SI) Fig. 8]. This finding suggests that fiber nucleation can be catalyzed by the surface of physical objects that can be eliminated by filtration.


Figure 1
View larger version (21K):
[in this window]
[in a new window]

  		Fig. 1. Kinetics of IAPP20-29 fiber formation. (A) Representative kinetics of 700 µM de novo reactions conducted in the presence (circles, red) and absence of a CH2Cl2:aqueous interface. Reactions without CH2Cl2 were prepared by diluting an organic stock solution of IAPP20-29 into an aqueous buffer solution at 25°C. The latter is prepared by diluting a filtered, 10x buffer solution with water either obtained directly from a Milli-Q purification system (triangles, green) or additionally passed through a 0.2-µm syringe filter (squares, blue). The t50 of these reactions are shown in Inset. (B) Kinetic profiles of reactions shown in A, with time axis renormalized to the t50 of the representative reactions. For triangles and squares, data are overlaid with fits to a sigmoid. A simulated reaction profile for actin (magenta) is shown as representative of wholly polymer-independent nucleation. The first 250 sec of a CH2Cl2:aqueous interface-mediated reaction profile fits to the function at2, where a is a constant (Inset, black line). (C) Measurement of reaction profiles at alternative protein concentrations. Shown are 2,000 µM (x) and 300 µM (+) reactions in the presence of a CH2Cl2:aqueous interface (Lower). Kinetics are also shown for 1 mM (squares) and 500 µM (circles) reactions without CH2Cl2 (Upper). After renormalizing time to the reaction t50, the respective profiles overlay (Right).

The effects of surface can be deliberately exaggerated. Reaction t50 are accelerated {approx}10-fold by the presence of a water:dichloromethane (CH2Cl2) interface. CH2Cl2 has low solubility in water and forms a separate liquid phase below the aqueous buffer. Reactions were initiated by first assembling an aqueous reaction, as described above, and then layering it on top of CH2Cl2. A 700 µM reaction has a t50 of 480 ± 170 sec in the presence and 3,400 ± 700 sec in the absence of this interface (Fig. 1A). Acceleration is attributable to this interface. Reactions conducted in the absence of an interface, but in the presence of buffer presaturated with CH2Cl2, have t50 decreased by only a factor of 1.9 ± 0.2 (SI Fig. 9). Furthermore, the extent of acceleration depends on the size of the interface relative to reaction volume (SI Fig. 10). Fibers formed in the presence and absence of CH2Cl2 are indistinguishable by negative stain transmission electron microscopy (TEM) (Fig. 2 A and B) and atomic force microscopy (SI Fig. 11). Furthermore, they have a common intersheet spacing (8.5 Å) by x-ray fiber diffraction (Fig. 2C) and seed fiber formation with the same efficiency (Fig. 2D). This suggests that the structure of fibers formed in the presence and absence of the CH2Cl2:aqueous interface are the same.


Figure 2
View larger version (26K):
[in this window]
[in a new window]

  		Fig. 2. Comparison of fibers formed in the presence and absence of a CH2Cl2:aqueous interface. (A and B) Negatively stained transmission electron micrographs of fibers formed at 1 mM in the absence (A) and presence (B) of a CH2Cl2:aqueous interface. (C) Representative x-ray diffraction patterns of unaligned fibers prepared in the presence (right) and absence (left) of the CH2Cl2:aqueous interface. Two reflections characteristic of beta-sheet structure are evident: {approx}4.7 Å, corresponding to the distance between peptides within the same beta-sheet, and {approx}8.5 Å, corresponding to the intersheet distance. (D) Fibers formed in the presence and absence of a CH2Cl2:aqueous interface seed fiber formation of IAPP20-29 monomer with the same efficiency. Representative reactions showing 800 µM IAPP20-29 de novo (squares) or seeded with 100 µM fibers formed in the presence (circles) and absence (triangles) of the interface. Lines shown are fits to sigmoidal and exponential equations, respectively.

In the presence of a CH2Cl2:aqueous interface, a single nucleation process is sufficient to describe the IAPP20-29 kinetic profile. To interpret assembly profiles, we, like others (1, 3–5, 7), first assume that the initial assembly steps are governed by unfavorable equilibria. The nucleus is defined here as the highest energy, least populated intermediate on the reaction pathway. The nucleus, and all states involved in its formation, are in equilibrium with monomer. To validate this assumption for IAPP20-29, we used 1H NMR and ultraviolet circular dichroism to determine that the protein is monomeric and has a random-coil structure during the lag phase (SI Figs. 12 and 13). Second, we assume that later steps in assembly are essentially irreversible. The first of these steps, which we term here as nucleation, is rate-limiting in the assembly pathway because it has the nucleus as a reactant. With these assumptions, the initial reaction profile is quadratic ([F](t) {propto} t2) (1), where t is time and [F](t) is the amount of polymerized material in monomer units. The early profile of a 700 µM IAPP20-29 reaction conducted in the presence of a CH2Cl2:aqueous interface fits clearly to this quadratic (Fig. 1B Inset). Furthermore, it is the interface that generates this curve shape; if CH2Cl2 is used only to saturate the aqueous buffer, then the initial time points are flat (SI Fig. 9). Actin is the canonical biopolymer that assembles via a single nucleation path (4). We note that the entire calculated profile of actin (Fig. 1B) is comparable to CH2Cl2-catalyzed assembly of 700 µM IAPP20-29. This is not unique to this protein concentration; reactions from 300 µM to 2 mM IAPP20-29 all have this time-renormalized profile (Fig. 1C and SI Fig. 14). Clearly, IAPP20-29 fiber assembly can be described by a single, fiber-independent nucleation mechanism in the presence of a CH2Cl2:aqueous interface.

A fiber-dependent mechanism of nucleation is evident in the absence of a CH2Cl2:aqueous interface. The initial portion of the IAPP20-29 reaction profile is flat and cannot be approximated by a quadratic (1) or higher order polynomial (3) (Fig. 1B). This property is evident when using detection methods that report on fiber growth [thioflavin T (ThT) and light scatter; SI Fig. 15] as well as methods sensitive to monomer loss (1H NMR; Fig. 3). In the latter study, the 1H NMR chemical shift dispersion is characteristic of random coil throughout the lag phase (SI Fig. 13). Furthermore, there are no time-dependent changes in chemical shift or linewidth that would be indicative of structure formation and/or oligomerization. Lastly, all of the protein is quantitatively present during the lag phase and diffuses as a single species (D = 3.0 ± 0.2 x 10–6 cm2/s) over an {approx}10-fold range of concentration (SI Fig. 13). This is within error of calculated estimates of D for random coil monomer: 2.8 x 10–6 cm2/s (22) or 4.1 ± 1.7 x 10–6 cm2/s (23), respectively. A reaction profile with a flat lag phase can be generated by a reaction that proceeds through a series of irreversible steps (i.e., downhill polymerization) (1). A flat lag phase results if the detection method is only sensitive to downstream states. However, because intermediates are not detectably populated by 1H NMR, such a mechanism is unlikely. In addition, a downhill polymerization model yields reaction kinetics that vary linearly with initial protein concentration (1). This is also not consistent with our observations reported below. A second possibility is that there are fiber-dependent nucleation processes. In this model, the probability that new fiber ends will form increases in proportion to fiber mass (8). Such a model is wholly consistent with our observations. Alone, this is not sufficient to describe the kinetic profile because fibers are not initially present in solution. Therefore, there must also be a contribution from a fiber-independent mechanism. Nevertheless, because the beginning of the reaction profile is completely flat, the vast majority of nucleation events must occur by the 2° mechanism. It is clear that both 1° and 2° nucleation are necessary to describe the entire reaction profile under wholly aqueous conditions.


Figure 3
View larger version (9K):
[in this window]
[in a new window]

  		Fig. 3. Kinetic profiles measured by light scatter and soluble monomer concentration are closely similar. A representative 700 µMIAPP20-29 fiber formation was split and monitored simultaneously by NMR (triangles) and by 90° light scatter at 400 nm (squares). Successive 1D 1H spectra were taken approximately every 10 min. The absolute monomer concentration was determined by comparing the area of the phenyl peak to the area of TMSP, which was added at a fixed concentration.

Fiber formation can be indirectly detected during the lag phase. A 1 mM reaction was diluted to 800 µM during the lag phase and compared with a reaction initiated at 800 µM. If fibers have formed during the lag phase of the former, then the diluted reaction will convert faster than a reaction initiated at 800 µM. Alternatively, if the lag phase is a measure of the time taken until the first 1° event, then the distribution of reaction t50 should be identical. Indeed, dilution to 800 µM at 150 sec gives t50 that are 1.36 ± 0.04-fold shorter than a reaction initiated at 800 µM (Fig. 4). As a control, a reaction initiated at 800 µM was agitated in an identical fashion at 150 sec. This gives t50 that are only 1.08 ± 0.09-fold accelerated. Importantly, the extent of acceleration depends on the time elapsed before dilution. Dilution at 300 sec gives t50 that are 1.65 ± 0.02-fold accelerated. Furthermore, reaction t50 show minimal (±7%) variation, indicating that stochastic occurrence of a 1° event is not governing the lag phase length (SI Fig. 16). Clearly, fiber formation has begun, albeit at low concentrations, during the lag phase.


Figure 4
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 4. Fiber formation occurs during the lag phase. A representative 1 mM reaction (circles, orange) was diluted to 800 µM at 150 sec (inverted triangles, magenta) and 300 sec (triangles, blue) into the lag phase with kinetics subsequently monitored. The ratio of t50 of diluted reactions to t50 of reactions initiated at 800 µM (x) were determined (Inset). The gray fraction shows the contribution of controls in which reactions initiated at 800 µM were agitated at 150 sec and 300 sec, respectively.

Fiber-independent nucleation has a reaction order of 4. For reactions with only 1° nucleation, the rate of fiber end formation is proportional to [A1]m, where A1 is free monomer and m is the reaction order. This gives t50 = {lambda}[Atot]–m/2, where Atot is the total protein and {lambda} is a constant (4, 6) (see SI Calculation 1A). Reactions with t50 varying over more than a 10-fold range were measured by varying protein concentration in the presence of a CH2Cl2:aqueous interface. The reaction is strongly concentration-dependent with a reaction order of {approx}4 (Fig. 5A). We note that in several analyses conducted on separate days, the reaction order was consistent but with varying {lambda}, presumably because of a systematic error associated with variations in exogenous surface. We accommodated this observation by performing a global analysis of 61 kinetic profiles on 8 separate days (Fig. 5B). Simultaneous fitting to a common reaction order yielded 3.9 ± 0.1 for the fiber-independent nucleation process.


Figure 5
View larger version (11K):
[in this window]
[in a new window]

  		Fig. 5. Determination of the reaction order for nucleation and elongation processes. (A) Representative data showing the concentration dependence of de novo reactions conducted in the presence (circles, red) and absence (triangles, green) of the CH2Cl2:aqueous interface. Reaction orders were obtained by simultaneous fitting of 8 and 13 independent data sets, respectively, to a power law with a common exponent. (B) The quality of the global analyses are shown by plotting measured vs. fitted t50 for all data sets. A line with a slope of one is shown in black. The reaction orders (Inset) are shown with errors corresponding to a 95% confidence interval. (C) Concentration dependence of fiber elongation. Reactions were conducted with concentrations of monomer ranging from 1 mM to 100 µM in the presence of 50 µM preformed fibers. Representative reaction profiles for two seeded kinetics containing 900 µM (triangles) and 400 µM (x) monomer are shown, respectively (Inset).

Fiber-dependent nucleation similarly has a reaction order of 4. The flat lag phase evident in profiles conducted in the absence of an interface suggests that 2° nucleation events predominate. Therefore, here but not in later analysis (see Discussion) we assume the rate of nucleus formation depends only on the amount of polymerized material, expressed as ([Atot] – [A1]). This gives a rate of fiber end formation proportional to [A1]n([Atot] – [A1]), resulting in t50 = {lambda}[Atot]–(n+1)/2, where n is the reaction order of 2° nucleation (see SI Calculation 2A). The one in the exponent of the latter accounts for the contribution of fiber surface to the apparent concentration dependence of the reaction t50. Any value of n ≥ 0 is allowable, with n = 0 corresponding to a fiber breakage mechanism. Reactions whose t50 span an {approx}100-fold range were measured by varying protein concentration (Fig. 5A). Above 3 mM, the t50 become as fast or faster than the mixing time (<1 min). Reactions at and below 500 µM (t50 > 2 h) show sufficient stochastic contributions that the data are not included in this analysis (SI Fig. 16). Global analysis of 112 reaction profiles collected over 13 separate days gives a reaction order of 4.0 ± 0.1 (Fig. 5B).

Elongation processes have a reaction order of 1. Fiber formation kinetics were conducted with 50 µM IAPP20-29 preformed fibers and soluble peptide concentrations ranging from 100 µM to 1 mM (Fig. 5C). The rate of elongation, as measured by the rate of monomer depletion, was taken from the slope of a line fit to the first {approx}400 sec of each reaction profile. If elongation of existing fiber ends occurs by addition of an oligomer of size l, then –d[A1]/dt = ke[E][A1]l, where E is fiber ends. Because the concentration of fiber ends is the same at the beginning of these reactions, a plot of log(d[A1]/dt) vs. log([Atot]) will have a slope of l. Analysis of three independent data sets give l = 0.90 ± 0.10, indicating that elongation proceeds via addition of monomer to fiber ends. Notably, this is distinct and smaller than the reaction orders obtained for nucleation shown above.

The enthalpic barrier to nucleation is the same in the presence and absence of a CH2Cl2:aqueous interface. This was assessed by measuring the temperature dependence of reaction t50. The t50 are exquisitely sensitive to temperature; for example, a 1 mM de novo reaction without a CH2Cl2:aqueous interface has a t50 of 1,400 ± 180 sec at 23°C and 5,300 ± 400 sec at 36.5°C. By 39°C the reaction is not observed to convert (t50 > 86,400 sec; data not shown). The Eyring equation enables us to relate the rate of nucleation to temperature, t50 {propto} e{Sigma}{Delta}Hi/2RT, where {Delta}Hi represents the enthalpic contribution from each of the barriers that determine the reaction t50 (this includes but is not limited to oligomerization equilibria governing the nucleus population, surface binding, and the barriers to nucleation and elongation; see SI Calculation 1B and SI Calculation 2B). Fits to this relation give {Sigma}{Delta}Hi = –36 ± 6 kcal/mol (Fig. 6). Similar time scales were achieved in the presence of a CH2Cl2:aqueous interface at 350 µM protein (Fig. 6 Inset). Remarkably, this yields a closely similar {Sigma}{Delta}Hi = –39 ± 3 kcal/mol. The contributions of some of the individual {Delta}Hi can be inferred. For example, elongation reactions can be conducted at temperatures above 39°C, giving rates that differ by <2 between 25°C and 45°C (SI Fig. 17). Similarly, the near-identical {Sigma}{Delta}Hi suggests that surface binding makes a comparatively small contribution. Therefore, the sensitivity of de novo reactions to temperature can be attributed predominantly to changes to oligomerization preequilibria and/or nucleation. Although we cannot yet make precise determinations of the individual {Delta}Hi, the fact that the {Sigma}{Delta}Hi are similar under markedly different reaction conditions strongly suggest that the two reactions are mechanistically similar.


Figure 6
View larger version (17K):
[in this window]
[in a new window]

  		Fig. 6. Temperature dependence of reactions conducted in the presence (circles, red) and absence (triangles, green) of the CH2Cl2:aqueous interface. Protein concentrations were 350 µM and 1 mM, respectively. Fits to determine activation enthalpies are shown (see text). Representative kinetics are shown (Inset).


   Discussion
Top
 Abstract
 Results
 Discussion
Materials and Methods
 Acknowledgements
 References
 
IAPP20-29 can form amyloid fibers by both a fiber-dependent and a fiber-independent nucleation mechanism. In the presence of a CH2Cl2:aqueous interface, fiber formation nucleates predominately by the 1° mechanism. Under wholly aqueous solution conditions, 1° nucleation is present but most nucleation occurs by the 2° mechanism. By making kinetic measurements under each of these solution conditions, we have shown that both nucleation processes can be described by a reaction order of 4 with net enthalpic contributions of approximately –37 kcal/mol to the apparent activation energy. Therefore, both 1° and 2° nucleation can be modeled as the same mechanism, differing only in the location of the event.

Global analysis of kinetics was used to verify our reaction orders without using our previous assumption that nucleation occurs by only a single mechanism under each solution condition (i.e., with or without the CH2Cl2:aqueous interface). Specifically, fiber formation kinetics were assessed for both conditions by using the following model:

Formula 1


Formula 2

where E represents fiber ends, A1 represents free monomer, (Atot – A1) represents fiber in monomer units, m, n, and l represent the reaction orders, and km, kn, and ke are effective rate constants associated with 1° nucleation, 2° nucleation, and elongation, respectively. Note that the amount of external surface present is reflected by the magnitude of km (SI Calculation 1A). Rate constants were fitted by numerical methods for various integer values of m and n (at l = 1) by simultaneously analyzing reaction profiles collected in the presence and absence of CH2Cl2 (Fig. 7A). Evaluation of {chi}2 as a function of m and n yields a best fit m = n = 4 (Fig. 7B). Thus, the same reaction orders are obtained whether or not we assume a single mechanism under each reaction condition. This does not represent a hidden property of the analysis, i.e., 1° does not mask 2° nucleation. To show this, Eqs. 1 and 2 were used to create synthetic data sets for alternative values of m and n. These were then subjected to the same global analysis and resulted in correct determination of m and n, for example m = 4, n = 2 (Fig. 7C). An important and generalizable observation (not shown) is that reaction profiles are concentration-dependent when m != n. Experimentally, IAPP20-29 shows no concentration dependence in its reaction profile (Fig. 1C and SI Fig. 14), consistent with m = n. Plainly, our global analysis faithfully extracts both 1° and 2° reaction orders.


Figure 7
View larger version (28K):
[in this window]
[in a new window]

  		Fig. 7. Global analysis of fiber formation. (A) Eqs. 1 and 2 were used to conduct a simultaneous fit of the concentration dependence of reaction profiles in the presence (circles, red) and absence (triangles, green) of the CH2Cl2:aqueous interface. For clarity, all data and the best fit (m = 4, n = 4, black) are shown with time renormalized to the t50 of an arbitrarily chosen data set. (B) {chi}2 for the fits in A are shown for fixed values of m and n. (C) Global analysis was also performed on synthetic data created with m = 4, n = 2 by using an equivalent number of data points as in B. Contours for B and C are shown on a log10 scale from lowest (blue) to highest (red) in increments of 0.33. (D and E) The rates of nucleation were calculated for a 700 µM profile by using the constants determined in A for aqueous (D) and CH2Cl2:aqueous (E) interface-mediated fiber formation, respectively. Loss of monomer A1(t) (green, –CH2Cl2, or red, +CH2Cl2), the rate of fiber-independent nucleation (dE1/dt, magenta), and the rate of fiber-dependent (dE2/dt, black) nucleation are shown. The point in time where the rates of nucleation by these mechanisms are equal is indicated with an arrow.

Interrogation of our global analysis enables us to assert the relative contribution of 1° and 2° nucleation at each time point during fibrillogenesis (Fig. 7 D and E). For interface-free reactions, 2° nucleation is dominant at all but the earliest times (Fig. 7D). Similarly, for reactions in the presence of CH2Cl2, the rate of 1° nucleation is dominant at all times before the t50 (Fig. 7E). This accounts for our ability to extract reaction order when assuming limiting conditions in the absence and presence of a CH2Cl2:aqueous interface (Fig. 5). However, the global analysis does not require the extremes of behavior provided by the interface. Alteration to the extent of 2° contributions provided, for example, by introduction of fiber as seed, would perturb the reaction profile and enable extraction of parameters. This should allow generalization of this approach, e.g., scission, as reported for Sup35 (13), is represented here by n = 0. Similarly, downhill polymerization, as reported for polyglutamine (7), is represented here by m = 1, kn = 0.

Surface-catalyzed nucleation is consistent with the high entropic cost of assembly. For nucleation to be unfavorable, the entropic contribution at a given temperature must exceed the enthalpic contribution of approximately –37 kcal/mol (Fig. 6). There are several ways that a surface can reduce the entropic penalty of assembly. First, there may be a high local concentration of peptides bound to the interface. Second, peptides may bind to the surface in an ordered, specific way that favors assembly. This is particularly true for parallel assemblies of beta-strands (24, 25). For IAPP20-29, acceleration is specific to certain interfaces. For example, CH2Cl2, chloroform, and phospholipid bilayers accelerate fiber formation and result in an apparent loss of secondary nucleation processes (SI Fig. 18). In contrast, hexanes, n-butanol, and carboxylate derivatized polystyrene microspheres do not have the effects reported here. Finally, surface catalysis could represent the stabilization of a high-energy nucleus by binding to the surface. This is analogous to lateral nucleation suggested for polymerization of hemoglobin S (26).

The extreme sensitivity of the IAPP20-29 fiber formation reaction to solution conditions can be explained if a surface, in addition to the protein itself, is required for nucleation. This result is not unique to IAPP20-29, because surfaces have been shown to strongly influence fiber formation in a number of other systems. Abeta, full-length IAPP, and {alpha}-synuclein fiber formation reactions can be manipulated by lipid membranes (27–30) and halogenated organic solvents including chloroform (31). Tau aggregation can also be triggered by surfaces (32, 33). In some cases, e.g., Abeta, interfaces can give rise to alternative forms of fibrillar and nonfibrillar aggregates as determined by the nature of the interface (34). Finally, assembly of hydrophobins into functional amyloid-like rodlets occurs at an air–water interface (35). Thus, the choice of conditions is critical for understanding assembly. Furthermore, it is likely that surfaces are of importance to in vivo fiber formation, because there are many surfaces with a rich variety of properties with which cellular proteins can interact.

In this work, we have equated 1° nucleation with 2° nucleation on a growing fiber. The kinetic profiles of fiber formation by full-length IAPP, Abeta, insulin, and the mammalian prion show that fiber-dependent nucleation plays a prominent role (9, 10, 12, 14). Similarly, the stochastic reaction kinetics reported for insulin, Abeta, and huntingtin can be rationalized with 2° nucleation (10, 14, 36). Finally, we note that imaging of both insulin (37) and IAPP (38) fiber formation suggests an intrinsic capacity of amyloid fibers to laterally stabilize their precursors. Although it is always possible to invoke an additional process to account for fiber-dependent reaction profiles, we have demonstrated here that this process need not be distinct from 1° nucleation.


   Materials and Methods
Top
 Abstract
 Results
 Discussion
 Materials and Methods
Acknowledgements
 References
 
Peptides. IAPP20-29 was synthesized in house and purified by RP-HPLC (see SI Materials and Methods).

Fiber Formation. IAPP20-29 stock solutions were prepared at 10 mM in DMSO. Fiber formation reactions were initiated by dilution into aqueous buffer (100 mM KCl/50 mM potassium phosphate, pH 7.4). Reaction buffer was augmented with DMSO to ensure consistent levels (10%) regardless of peptide concentration. Fiber formation reactions were monitored either by 90° light scatter or fluorescence enhancement of thioflavin T (ThT), added at 10 µM. Transmission electron micrograph and x-ray diffraction of fibers were conducted on samples from 1 mM reactions. For additional details, see SI Materials and Methods.

Data Analysis. The t50 of reactions were determined by finding the time corresponding to the half-maximum signal for each kinetic profile. All reported errors reflect one standard deviation of at least three separate measurements. Alternatively, confidence intervals were determined by Monte Carlo methods and reflect ±95% of the population of fitted parameters. Fitting, global analysis, and confidence intervals were determined by using MATLAB (MathWorks). For additional details, see SI Materials and Methods.


   Acknowledgements
Top
 Abstract
 Results
 Discussion
 Materials and Methods
 Acknowledgements
References
 
We thank Drs J. Knight and E. Rhoades for critical reading of the manuscript and J. Williamson (NMR), M. Calabrese (x-ray), B. Koo (electron microscopy), B. Piekos (electron microscopy), V. Unger (electron microscopy), Y. Kwon (atomic force microscopy), P. Sung (atomic force microscopy), and J. Hebda (liposomes) for assistance, training, and/or use of indicated tools used in this work. A.M.R. is a Howard Hughes Medical Institute Predoctoral Fellow. This work was supported by National Institutes of Health Grant DK54899.


   Footnotes
 

Abbreviations: IAPP, islet amyloid polypeptide.

{ddagger}To whom correspondence should be addressed. E-mail: andrew.miranker{at}yale.edu

Author contributions: A.M.R. and A.D.M. designed research; A.M.R. performed research; A.M.R. and A.D.M. analyzed data; and A.M.R. and A.D.M. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

This article contains supporting information online at www.pnas.org/cgi/content/full/0703306104/DC1.

© 2007 by The National Academy of Sciences of the USA


   References
Top
 Abstract
 Results
 Discussion
 Materials and Methods
 Acknowledgements
 References
 

  1. Ferrone, F. (1999) Methods Enzymol 309, 256–274.[ISI][Medline]
  2. Rochet, JC & Lansbury, PT. (2000) Curr Opin Struct Biol 10, 60–68.[CrossRef][ISI][Medline]
  3. Flyvbjerg, H, Jobs, E & Leibler, S. (1996) Proc Natl Acad Sci USA 93, 5975–5979.[Abstract/Free Full Text]
  4. Oosawa, F & Asakura, S. eds. Thermodynamics of the Polymerization of Protein (1975) (Academic, London,).
  5. Powers, ET & Powers, DL. (2006) Biophys J 91, 122–132.[CrossRef][ISI][Medline]
  6. Goldstein, RF & Stryer, L. (1986) Biophys J 50, 583–599.[ISI][Medline]
  7. Chen, S, Ferrone, FA & Wetzel, R. (2002) Proc Natl Acad Sci USA 99, 11884–11889.[Abstract/Free Full Text]
  8. Ferrone, FA, Hofrichter, J & Eaton, WA. (1985) J Mol Biol 183, 611–631.[CrossRef][ISI][Medline]
  9. Padrick, SB & Miranker, AD. (2002) Biochemistry 41, 4694–4703.[CrossRef][Medline]
  10. Hortschansky, P, Schroeckh, V, Christopeit, T, Zandomeneghi, G & Fändrich, M. (2005) Protein Sci 14, 1753–1759.[Abstract/Free Full Text]
  11. Hall, D & Edskes, H. (2004) J Mol Biol 336, 775–786.[CrossRef][ISI][Medline]
  12. Collins, SR, Douglass, A, Vale, RD & Weissman, JS. (2004) PLoS Biol 2, e321.[CrossRef][Medline]
  13. Tanaka, M, Collins, SR, Toyama, BH & Weissman, JS. (2006) Nature 442, 585–589.[CrossRef][Medline]
  14. Manno, M, Craparo, EF, Martorana, V, Bulone, D & San Biagio, PL. (2006) Biophys J 90, 4585–4591.[CrossRef][ISI][Medline]
  15. Smith, JF, Knowles, TP, Dobson, CM, Macphee, CE & Welland, ME. (2006) Proc Natl Acad Sci USA 103, 15806–15811.[Abstract/Free Full Text]
  16. Hull, RL, Westermark, GT, Westermark, P & Kahn, SE. (2004) J Clin Endocrinol Metab 89, 3629–3643.[Abstract/Free Full Text]
  17. Ritzel, RA, Meier, JJ, Lin, CY, Veldhuis, JD & Butler, PC. (2007) Diabetes 56, 65–71.[Abstract/Free Full Text]
  18. Ashburn, TT, Auger, M & Lansbury, PT Jr. (1992) J Am Chem Soc 114, 790–791.[CrossRef][ISI]
  19. Rhoades, E, Agarwal, J & Gafni, A. (2000) Biochim Biophys Acta 1476, 230–238.[CrossRef][Medline]
  20. Jaikaran, ET & Clark, A. (2001) Biochim Biophys Acta 1537, 179–203.[Medline]
  21. LeVine, H. (1993) Protein Sci 2, 404–410.[Abstract]
  22. Danielsson, J, Andersson, A, Jarvet, J & Gräslund, A. (2006) Magn Reson Chem 44, S114–S121.[CrossRef][ISI][Medline]
  23. Kohn, JE, Millett, IS, Jacob, J, Zagrovic, B, Dillon, TM, Cingel, N, Dothager, RS, Seifert, S, Thiyagarajan, P & Sosnick, TR, et al. (2004) Proc Natl Acad Sci USA 101, 12491–12496.[Abstract/Free Full Text]
  24. Xu, Z, Brauner, JW, Flach, CR & Mendelsohn, R. (2004) Langmuir 20, 3730–3733.[CrossRef][ISI][Medline]
  25. Kerth, A, Erbe, A, Dathe, M & Blume, A. (2004) Biophys J 86, 3750–3758.[CrossRef][ISI][Medline]
  26. Rotter, MA, Kwong, S, Briehl, RW & Ferrone, FA. (2005) Biophys J 89, 2677–2684.[CrossRef][ISI][Medline]
  27. Knight, JD & Miranker, AD. (2004) J Mol Biol 341, 1175–1187.[CrossRef][ISI][Medline]
  28. Gorbenko, GP & Kinnunen, PK. (2006) Chem Phys Lipids 141, 72–82.[CrossRef][ISI][Medline]
  29. Davidson, WS, Jonas, A, Clayton, DF & George, JM. (1998) J Biol Chem 273, 9443–9449.[Abstract/Free Full Text]
  30. Rhoades, E, Ramlall, TF, Webb, WW & Eliezer, D. (2006) Biophys J 90, 4692–4700.[CrossRef][ISI][Medline]
  31. Nichols, MR, Moss, MA, Reed, DK, Hoh, JH & Rosenberry, TL. (2005) Biochemistry 44, 165–173.[CrossRef][Medline]
  32. Chirita, CN, Congdon, EE, Yin, H & Kuret, J. (2005) Biochemistry 44, 5862–5872.[CrossRef][Medline]
  33. Kuret, J, Congdon, EE, Li, G, Yin, H, Yu, X & Zhong, Q. (2005) Microsc Res Tech 67, 141–155.[CrossRef][ISI][Medline]
  34. Kowalewski, T & Holtzman, DM. (1999) Proc Natl Acad Sci USA 96, 3688–3693.[Abstract/Free Full Text]
  35. Mackay, JP, Matthews, JM, Winefield, RD, Mackay, LG, Haverkamp, RG & Templeton, MD. (2001) Structure(London) 9, 83–91.
  36. Colby, DW, Cassady, JP, Lin, GC, Ingram, VM & Wittrup, KD. (2006) Nat Chem Biol 2, 319–323.[CrossRef][ISI][Medline]
  37. Jansen, R, Dzwolak, W & Winter, R. (2005) Biophys J 88, 1344–1353.[CrossRef][ISI][Medline]
  38. Goldsbury, C, Kistler, J, Aebi, U, Arvinte, T & Cooper, GJ. (1999) J Mol Biol 285, 33–39.[CrossRef][ISI][Medline]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg    What's this?


This article has been cited by other articles in HighWire Press-hosted journals:


Home page
 Proc. Natl. Acad. Sci. USAHome page
M. F. M. Engel, L. Khemtemourian, C. C. Kleijer, H. J. D. Meeldijk, J. Jacobs, A. J. Verkleij, B. de Kruijff, J. A. Killian, and J. W. M. Hoppener
Membrane damage by human islet amyloid polypeptide through fibril growth at the membrane
PNAS, April 22, 2008; 105(16): 6033 - 6038.
[Abstract] [Full Text] [PDF]

Home page
 Protein Eng Des SelHome page
B. W. Koo, J. A. Hebda, and A. D. Miranker
Amide inequivalence in the fibrillar assembly of islet amyloid polypeptide
Protein Eng. Des. Sel., March 1, 2008; 21(3): 147 - 154.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Figures Only
Right arrow Full Text (PDF)
Right arrow Supporting Information
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a colleague
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My File Cabinet
Right arrow Download to citation manager
Right arrow Request Copyright Permission
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via CrossRef
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ruschak, A. M.
Right arrow Articles by Miranker, A. D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ruschak, A. M.
Right arrow Articles by Miranker, A. D.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg  
What's this?