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Published online before print April 5, 2006, 10.1110/ps.051803606
Protein Science (2006), 15:1162-1174. Published by Cold Spring Harbor Laboratory Press. Copyright © 2006 The Protein Society
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Secondary structure and dynamics of micelle bound beta- and {gamma}-synuclein

Yoon-hui Sung and David Eliezer

Department of Biochemistry and Program in Structural Biology, Weill Medical College of Cornell University, New York, New York 10021, USA

(RECEIVED August 23, 2005; FINAL REVISION January 27, 2006; ACCEPTED January 28, 2006)


   Abstract
TOP
 Abstract
Introduction
 Results
 Discussion
 Conclusions
 Materials and methods
 Electronic supplemental material
 References
 
We have used solution state NMR spectroscopy to characterize the secondary structure and backbone dynamics of the proteins beta- and {gamma}-synuclein in their detergent micelle-bound conformations. Comparison of the results with those previously obtained for the Parkinson's disease-linked protein {alpha}-synuclein shows that structural differences between the three homologous synuclein family members are directly related to variations in their primary amino acid sequences. An 11-residue deletion in the lipid-binding domain of beta-synuclein leads to the destabilization of an entire segment of the micelle-bound helical structure containing the deletion site. The acidic C-terminal tail region of {gamma}-synuclein, which displays extensive sequence divergence, is more highly disordered than the corresponding regions in the other two family members. The observed structural differences are likely to mediate functional variations between the three proteins, with differences between {alpha}- and beta-synuclein expected to revolve around their lipid interactions, while differences in {gamma}-synuclein function are expected to result from different protein–protein interactions mediated by its unique C-terminal tail.

Keywords: synuclein; Parkinson's; membrane proteins; amyloid; protein aggregation


   Introduction
TOP
 Abstract
 Introduction
Results
 Discussion
 Conclusions
 Materials and methods
 Electronic supplemental material
 References
 
The synucleins are a small, highly conserved family of vertebrate proteins that includes {alpha}-synuclein (aS), beta-synuclein (bS), and {gamma}-synuclein (gS) (George 2002). aS and bS are expressed primarily in the central nervous system (Nakajo et al. 1993; Ueda et al. 1993; Jakes et al. 1994) with a notable localization to presynaptic terminals (Nakajo et al. 1994; Iwai et al. 1995). gS expression was initially reported in the peripheral nervous system (Akopian and Wood 1995) and breast cancer tissue (Ji et al. 1997), but gS is also expressed in the brain (Buchman et al. 1998; Lavedan et al. 1998), including the substantia nigra (Lavedan et al. 1998; Brenz Verca et al. 2003). The normal physiological roles of synuclein proteins remain poorly understood, although accumulating evidence implicates aS, the best characterized member of the family, in the regulation of synaptic vesicle function (Abeliovich et al. 2000; Murphy et al. 2000; Cabin et al. 2002; Schluter et al. 2003) and in the homeostasis and trafficking of the neurotransmitter dopamine (Perez and Hastings 2004; Sidhu et al. 2004; Yu et al. 2005).

Several observations have supported a role for aS in the pathogenesis of Parkinson's disease (PD). aS has been identified as a primary protein component of Lewy body and Lewy neurite deposits in PD and related disorders, where it is found in the form of amyloid fibril aggregates (Spillantini et al. 1998). Moreover, missense mutations (A53T, A30P, and E46K) (Polymeropoulos et al. 1997; Kruger et al. 1998; Zarranz et al. 2004) and gene triplication (Bradbury 2003; Singleton et al. 2003; Farrer et al. 2004) of aS have been shown to be genetically linked to early-onset familial parkinsonism. Although bS and gS have not been found in Lewy bodies or Lewy neurites, both proteins have been reported to be associated with hipppocampal axon pathology in PD (Galvin et al. 1999) as well as with deposits in other neurodegenerative disorders (Galvin et al. 2000).

In addition to their overlapping expression, all synuclein proteins show a high level of sequence homology with each other (Fig. 1). Each of the three family members is composed of an N-terminal lipid-binding domain, containing a series of 11-residue imperfect repeats, and an acidic C-terminal domain. Among the human family members, aS is 62% identical and 79% homologous to bS, aS is 50% identical and 74% homologous to gS, and bS is 47% identical and 66% homologous to gS. The bS N-terminal region lacks 11 central hydrophobic residues that correspond to parts of repeats 6 and 7 of aS and gS. Both aS and bS contain two 16-residue tandem repeats in their acidic C-terminal domain (Nielsen et al. 2001), while gS lacks these repeats and is highly divergent from the other family members in this region, maintaining only its highly acidic character. The highly conserved N-terminal region is known to be important for the lipid interactions of the synucleins (Perrin et al. 2000; Eliezer et al. 2001) and the highly acidic C-terminal region has been suggested to possess chaperone-like activity (Kim et al. 2002), to regulate the aggregation of aS (Murray et al. 2003) and to mediate protein–protein interactions (Jensen et al. 1999; Eliezer et al. 2001; Payton et al. 2004).


Figure 1
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  		Figure 1. Sequence alignment of human aS, bS, and gS. The imperfect 11-residue repeats are delineated by spaces. The KTKEGV consensus sequence within each repeat is underlined. Boldface characters indicate a difference from the aS sequence.

 
In vitro, aS has been demonstrated to adopt either a highly unstructured free state (Weinreb et al. 1996; Eliezer et al. 2001) or a highly helical lipid-bound state (Davidson et al. 1998; Eliezer et al. 2001). It is likely that both conformations observed in vitro occur in vivo as well, with the lipid-bound state representing the membrane- or synaptic vesicle-associated protein fraction and the free state representing the cytosolic fraction. Because the normal function of aS appears to involve the regulation of synaptic vesicles and because a number of purported aS-interacting proteins are membrane associated, the lipid-bound conformation of the protein is thought to be crucial for carrying out its normal function. Detailed NMR studies of aS bound to lipid-mimicking detergent micelles have revealed that the N-terminal lipid-binding domain adopts two separate amphipathic helices that reside on the micelle surface in a manner similar to that anticipated for apolipoproteins containing similar 11-residue repeats (Bussell and Eliezer 2003; Chandra et al. 2003; Bisaglia et al. 2005; Bussell et al. 2005; Ulmer et al. 2005). Although the hinge region between the two helices probably adopts a well-defined structure, the two helices do not appear to interact with one another (Bussell et al. 2005; Ulmer et al. 2005). Corresponding ESR studies of lipid vesicle-bound aS support these observations (Jao et al. 2004).

In contrast to aS, very little information is available regarding the structural properties of bS and gS. Spectroscopic studies of both proteins have confirmed that their free states behave similarly to that of aS, although bS may be somewhat more highly unfolded (Uversky et al. 2002). The very high degree of conservation in the lipid-binding N-terminal domains of all three synucleins strongly suggests that both bS and gS will, like aS, bind to lipid membranes and adopt a highly helical structure. As of yet, however, there is no reported data on the conformations of membrane-bound bS and gS. Here we present a characterization of the structural and dynamic properties of detergent micelle-bound bS and gS and compare them with those of aS.


   Results
TOP
 Abstract
 Introduction
 Results
Discussion
 Conclusions
 Materials and methods
 Electronic supplemental material
 References
 
Circular dichroism
Figure 2 shows the far-UV circular dichroism (CD) spectra of bS and gS bound to SDS detergent micelles, with the corresponding spectrum of aS included for comparison. The spectra of aS and gS are almost indistinguishable, while that of bS suggests a slightly less helical structure, as indicated by the decreased signal at 222 nm and a slight shift of the minimum at 208 nm toward shorter wave lengths. Similar results were obtained for both proteins in the presence of either lysophospholipid micelles or small unilamellar phospholipid vesicles (Supplemental Figure S1), indicating that the conformations of the lipid-bound proteins closely resemble those of the detergent micelle-bound states.


Figure 2
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  		Figure 2. Far-UV CD spectra of SDS micelle-bound synuclein family members aS ({bigtriangleup}), bS ({square}), and gS ({circ}) at pH 7.4.

 
Chemical shifts
To compare the secondary structure content of micelle-bound bS and gS with aS, the NMR backbone and Cbeta resonances were assigned for both proteins. Proton–nitrogen correlation spectra of micelle-bound bS and gS (Fig. 3) show a degree of resonance dispersion that is higher than that observed in equivalent spectra of the free proteins (data not shown), and is consistent with the presence of well-formed elements of secondary structure, but not with the extensive tertiary interactions evident in well-folded globular proteins. To identify the specific residues involved in secondary structures, the chemical shifts of the assigned C{alpha}, CO, Cbeta, and H{alpha} resonances for both bS and gS were compared to the values that would be expected for a random coil conformation. The differences between the observed and random coil chemical shifts, known as secondary shifts, have been shown empirically to correlate closely with the presence of secondary structure in proteins (Wishart and Case 2002). Helical structure leads to positive C{alpha} and CO secondary shifts and negative H{alpha} and Cbeta secondary shifts, while extended or strand-like structure leads to negative C{alpha} and CO and positive H{alpha} and Cbeta secondary shifts. Unstructured flexible backbone regions exhibit secondary shifts closer to zero values. The reliability of these secondary shifts values is in the order C{alpha}, CO, H{alpha}, Cbeta to identify helical propensity, and in the order H{alpha}, Cbeta, C{alpha}, CO to identify strand propensity (Yao et al. 1997; Wishart and Case 2002).


Figure 3
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  		Figure 3. 1H-15N proton–nitrogen correlation spectra of bS (A) and gS (B) in the presence of SDS micelles at pH 7.4. Sequence-specific resonance assignments are indicated. Unlabeled peaks correspond to side-chain NH2 groups.

 
Figure 4 shows the secondary chemical shifts of all four of these nuclei as a function of residue number for both bS and gS. The overall patterns of the secondary chemical shifts of bS and gS are quite similar to previous observations for aS, showing the extensive helical propensity of the N-terminal lipid-binding domain and the relatively unstructured nature of acidic C-terminal tail. The C{alpha} shifts suggest that the helical structure ends around position 83 for bS and 94 for gS, consistent with the 11 residues missing from the lipid-binding domain of bS. The remaining secondary shifts are not in exact agreement regarding the endpoints inferred from C{alpha} shifts, but clearly indicate an end to the helical structure within a few residues of these locations. The C{alpha} secondary shifts from bS also show clear evidence for a break in the helical structure of the N-terminal domain at residues 43 and 44, where significantly lower values are observed. Similar behavior was previously observed for aS. In the case of gS, a slightly smaller C{alpha} secondary shift was observed for residue 43, but the decrease relative to the surrounding residues is not dramatic and would not by itself indicate a clear break in the helical structure. Nevertheless, for both bS and gS, the secondary chemical shifts of the other nuclei strongly support a break in the helical structure of the N-terminal domains, as indicated by small or negative CO secondary shifts at residues 36 and 41, small or positive H{alpha} secondary shifts at residues 36, 42, and 43, and small or positive Cbeta secondary shifts at residues 38 through 44.


Figure 4
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  		Figure 4. Deviation of C{alpha}, CO, H{alpha}, and Cbeta chemical shifts from random coil values, for bS (A) and gS (B) in the presence of SDS micelles at pH 7.4.

 
In addition to indicating a break in the helical N-terminal region, the secondary shifts for both proteins also suggest a lower degree of helical structure both at the N-terminal and C-terminal regions of the helical domain. The first 11 residues exhibit lower amplitude C{alpha} and H{alpha} shifts and mixed CO and Cbeta secondary shifts for both proteins. The last seven to eight residues (77–83 for bS and 87–94 for gS) also show low and decreasing C{alpha} and H{alpha} shifts and mixed CO and Cbeta shifts. Finally, for bS, it appears that the residues 65–76 are also only marginally helical, with relatively small (positive) C{alpha} shifts and mixed CO, H{alpha}, and Cbeta shifts. The gS data also indicate a decrease in helical structure content around position 65, but this appears to be confined to only a few residues, and robust evidence for helix resumes at position 67. The latter feature was also observed for aS.

Nuclear Overhauser Effects (NOEs)
In addition to chemical shifts, we examined Nuclear Overhauser Effects (NOEs) between sequential amide protons in micelle-bound bS and gS. The distance that these NOEs report on is short in helical conformations and considerably longer in extended or strand structure. Both bS and gS exhibit strong HNHN NOEs (Fig. 5) in the regions identified as helical through the chemical shift analysis, but show very few NOEs in the C-terminal region, consistent with a lack of well-defined structure in the acidic tail regions. For both proteins, however, NOEs of significant intensity continue beyond the termination point of helical structure, as inferred from secondary chemical shifts. For bS, strong NOEs continue until position 92 and for gS until position 101. These NOEs suggest that a short stretch (7–9 residues) of defined, locally compact structure exists in the lipid-binding domains of both proteins just C-terminal to the end of helical structure. This is again similar to observations made for aS.


Figure 5
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  		Figure 5. Sequential amide proton to amide proton NOEs in bS (top panel) and gS (bottom panel) in the presence of SDS micelles at pH 7.4. The average of the forward and backward NOEs is shown. Strong NOEs are associated with short NH–NH distances such as those present in helical structure. The letters XX indicate resonance overlap between neighboring residues, precluding a determination of the presence or absence of these sequential NOEs. Absent NOEs between proline residues, which lack an amide proton, and both preceding and subsequent residues are indicated by filled circles.

 
Both sets of NOE data also show significantly lower NOE intensities between residues 42 and 43, consistent with the secondary shift indications for a break in the helical structure around this location. A weak NOE is also observed between residues 64 and 65 for bS. This could be considered consistent with the reduced indication for helical structure beyond this position in the secondary chemical shifts of bS. Surprisingly, however, strong NOEs resume at position 66 and extend until the end of the lipid-binding domain. NOE measurements preferentially detect conformations where internuclear distances are short. Therefore, it seems likely that for bS, the region C-terminal to residue 65 (residues 65–83) is dynamic, populating a helical structure some fraction of the time, as reflected in the strong NHNH NOEs and also in the C{alpha} secondary shifts, but also another, presumably unstructured, conformation. Supporting this idea is a notable increase in NOE intensity after position 65. This intensity increase is associated with a concomitant increase in the amplitude of the corresponding diagonal resonances (data not shown), suggesting that it originates from an increase in the overall mobility of this region rather than from shorter distances between successive amide protons.

A number of weaker NOEs are also evident in the data from gS, at positions 59–60 and 78–79, for example. Neither of these is correlated with a clear feature in the chemical shift data, suggesting that they do not represent major deviations from helical structure. Starting around position 83, however, a similar increase in NOE intensity to that observed past position 64 in bS is observed, suggesting that here, too, the C-terminal end of the helical lipid-bound structure may be somewhat dynamic.

Scalar coupling constants
Vicinal coupling constants, 3JHNH{alpha} are sensitive to the backbone torsion angle {varphi}, and can provide an additional indication of secondary structure. In well-defined helical regions, 3JHNH{alpha} values of around 4 Hz are expected, whereas in well-formed beta-strands, values of ~9 Hz are typical (Wuthrich 1986). 3JHNH{alpha} values for bs and gS are shown in Figure 6. Although the data are quite noisy, for bS smaller values consistent with helical structure are observed from residues 1 to 35, and again from residues 45 to 64. Residues 36–44 show higher values, consistent again with a break in helical structure in this region. Residues 65–94 also show higher values, with a short dip around positions 78 and 79. This is consistent with the secondary chemical shifts and indicates that any helical structure formed in this region is likely to be relatively unstable and possibly dynamic. Beyond position 94 a further slight increase in the coupling constants is consistent with an extended flexibly C-terminal tail.


Figure 6
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  		Figure 6. 3JHNH{alpha} coupling constants for bS (top panel) and gS (bottom panel) obtained from HNHA spectra in the presence of SDS micelles at pH 7.4. Bars in gray indicate data points that may contain contributions from overlapping resonances. Asterisks indicate proline residues, which lack an amide proton.

 
In the case of gS, the 3JHNH{alpha} coupling constants appear to be somewhat higher in general for the lipid-bound helical region. The values in the N-terminal region prior to position 36 are not as low as those observed for bS, especially for the first 10 residues, which also show only weakly helical structure in the chemical shift data as well as an increase in NOE intensity as described above. Similarly, positions 36–44 also show higher coupling constant values, consistent with an interruption of helical structure, and there is a gradual increase from positions 84 to 93, beyond which the high values expected for the flexible tail region are observed.

Dynamics
To investigate the picosecond–nanosecond time scale backbone dynamics of micelle-bound bS and gS, we measured the 15N longitudinal and transverse relaxation rate constants R1 and R2, as well as the heteronuclear 15N-(1H) steady-state NOE. In the extreme narrowing limit (motions occurring much faster than the characteristic NMR time scale 1/{omega}0) R1 and R2 take on the same value. For motions occurring on time scales similar to or slower than 1/{omega}0, increases in R1 and decreases in R2 suggest relatively faster motions. Increases in R2 can also reflect the presence of slower (millisecond–microsecond) motions associated with chemical exchange processes. The heteronuclear NOE is a sensitive indicator of fast motions arising from local flexibility, with positive values near 1 indicating the least flexible regions, and smaller or negative values indicating greater flexibility.

Inspection of the relaxation parameters obtained for bS and gS (Fig. 7) clearly shows a demarcation between the micelle-associated N-terminal domain and the flexible C-terminal tails for both proteins. For bS, the heteronuclear NOE values are negative beginning with position 98, although many values near zero, as well as two clearly negative values are apparent at positions 89–97 as well. Positions 89–97 also show distinctly lower values of R2 than the remainder of the N-terminal domain, but these values decrease even further past position 97. For gS, the heteronuclear NOEs turn slightly negative at position 102, with a further significant decrease at position 107. R2 values decrease slightly around position 93, then more dramatically from positions 101–107.


Figure 7
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  		Figure 7. R1, R2, and 15N-(1H) steady-state NOE relaxation parameters for backbone 15N nuclei in bS (A) and gS (B) in the presence of SDS micelles at pH 7.4.

 
Within the N-terminal domains of each protein more subtle variations are apparent. Both proteins show distinctly lower R2 values for positions 1–7 and a slight dip in the R2 data for residues ~30–40. For bS, residues 67–88 show clearly lower R2 and heteronuclear NOE values, consistent with indications of lower helicity from chemical shifts and coupling constants and the greater mobility suggested by the NOE data for these regions. For gS, there is a narrower depression in the R2 data at position 65, where again chemical shifts suggest an interruption of helical structure.

Within the C-terminal domains of both proteins, past positions 97 for bS and 107 for gS, the R2 values are similar in amplitude to the R1 values, implying an approach to the extreme narrowing limit, in agreement with the large amplitude negative heteronuclear NOE values.

The three relaxation parameters were also analyzed using the model free formalism (Lipari and Szabo 1982a,b) in order to generate the order parameter S2, which describes the amplitude of fast motions experienced by the individual NH bond vectors (Fig. 8). High S2 values, approaching 1, indicate limited motions and greater rigidity, whereas lower values indicate increasingly larger amplitude motions and greater flexibility. The S2 data reinforce the features observed in the raw relaxation parameters. The transition to the disordered tail can be seen clearly at positions 88 and 101 for bS and gS, respectively. For bS, lower S2 values are clearly evident from positions 67 to 88, while for gS, a slight dip in S2 is seen from positions 30–40 and also at position 65.


Figure 8
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  		Figure 8. The generalized order parameter S2 for bS (top panel) and gS (bottom panel) in the presence of SDS micelles at pH 7.4. No data are shown for proline residues, which lack an amide proton, for residues with overlapped signals, or for gS residue 18, which was not well fit.

 

   Discussion
TOP
 Abstract
 Introduction
 Results
 Discussion
Conclusions
 Materials and methods
 Electronic supplemental material
 References
 
The synuclein family gained attention through the discovery that aS is genetically linked to familial PD through either missense mutations (Polymeropoulos et al. 1997; Kruger et al. 1998; Zarranz et al. 2004) or gene triplication (Bradbury 2003; Singleton et al. 2003; Farrer et al. 2004) and is also phenomenologically linked to sporadic PD through the presence of aS amyloid fibril aggregates in the hallmark Lewy body deposits associated with the disease (Spillantini et al. 1998). However, the exact mechanism by which aS is involved in the pathogenesis of PD remains unclear. Many believe that aS aggregation is key in mediating the toxic effects of the protein through the action of either the mature amyloid fibrils or some oligomeric species formed during fibril assembly. This view is supported by a variety of results, including the facts that PD-linked mutations enhance aS oligomerization in vitro (Conway et al. 1998, 2000; Narhi et al. 1999), by the fact that increased levels of aS can cause disease in humans (Bradbury 2003; Singleton et al. 2003; Farrer et al. 2004) or in animal models (Feany and Bender 2000; Masliah et al. 2000), and that amyloid fibrils in general are toxic to cells (Bucciantini et al. 2002). An alternative possibility is that a deficiency in the normal function of aS is responsible for its role in PD. This viewpoint is supported by the facts that aS function appears to revolve around dopamine trafficking and homeostasis and that PD is fundamentally a dopamine deficit disorder (Perez and Hastings 2004; Sidhu et al. 2004; Yu et al. 2005). The two possibilities are not necessarily mutually exclusive, and may even be linked, as enhanced aggregation of aS may serve to deplete the functional pool of the protein.

The other members of the synuclein family have been largely ignored until recently because of the absence of any genetic link to PD. Nevertheless, a number of recent reports suggest that bS and possibly gS as well may influence the role of aS in PD by influencing its aggregation (Hashimoto et al. 2001; Uversky et al. 2002; Park and Lansbury 2003), and may even play a direct role themselves in neurodegenerative processes (Galvin et al. 1999, 2000). It also appears that the different synucleins may have overlapping functions, as a recent aS/bS double-knockout mouse has a more severe phenotype than is obtained by knocking out aS alone (Chandra et al. 2004). A similarity of function between all three members of the synuclein family is also indicated by the high degree of homology between the three proteins, particularly in their lipid-binding N-terminal domains. Nevertheless, differences in the sequences of the three proteins in both their N-terminal and C-terminal domains must be responsible for those differences that do exist in their individual functions, as well as for their different roles in disease. In order to clarify both the similarities and the differences between the three synuclein family members at a structural level, we have characterized the folded functional forms of bS and gS bound to lipid-mimetic detergent micelles using NMR, in a manner similar to that employed in our own and others’ recent studies of aS (Eliezer et al. 2001; Bussell and Eliezer 2003, 2004; Chandra et al. 2003; Bisaglia et al. 2005; Bussell et al. 2005; Ulmer et al. 2005).

Domain boundaries
As expected based on the sequence alignment shown in Figure 1, all of the data presented here indicate that bS and gS, like aS, consist of a helical lipid-binding N-terminal domain containing characteristic 11-residue pseudo-repeats and an acidic C-terminal tail that does not associate tightly with membranes. The endpoint of the helical structure of the N-terminal domain is essentially the same for gS as for aS, occurring at position 94, a few residues past the end of the seventh and last 11-mer repeat, as indicated primarily by chemical shifts. For bS, which is missing 11 residues from the N-terminal domain, the end of the helical structure is observed at residue 83. For both proteins, there is a clear indication that several residues beyond the end of helical structure retain some degree of ordered structure, as indicated by the continuation of strong NHNH NOEs and restricted backbone motions evident in the backbone dynamics data. For gS, this region extends approximately to residue 101 and again exactly parallels the behavior of aS, where a similar pattern of NOEs (Bussell and Eliezer 2003) and dynamics data (Bisaglia et al. 2005; Bussell et al. 2005; Ulmer et al. 2005) can be seen, and for which a recent molecular fragment replacement-based structure calculation confirmed an ordered but extended structure (Ulmer et al. 2005). For bS, this region extends to residue 89. Interestingly, the backbone motions of residues 89–97 for bS and 102–107 for gS are also somewhat restricted, and it appears that these residues may act as linker regions between the micelle-associated N-terminal domains and the free and flexible C-terminal domains. Residues within the linkers are affected by the overall dynamics of both domains, whereas residues on either side are effectively isolated from the dynamics of the other domain.

Helix break
The helical structure of the N-terminal domain of both bS and gS is clearly interrupted around position 42 in the same manner previously observed for aS (Bussell and Eliezer 2003; Chandra et al. 2003), as indicated by both chemical shifts and NOEs. Although there are several amino acid substitutions around this region in bS and gS (K45R in bS; L38M and S42A in gS), they are all reasonably conservative, consistent with the region behaving similarly in all three proteins. Notably Y39, the only tyrosine residue found within the N-terminal lipid-binding region, is conserved in all three proteins. The unique combination of both an aromatic and a polar character of tyrosine (tryptophan has this property as well but does not appear in any synuclein sequence) makes it a common membrane-interface residue, and may be crucial for causing the interruption of the helical structure of all three synuclein proteins (Bussell and Eliezer 2003). The break in the helical structure of aS leads to a topology in which the two separate helices are oriented antiparallel to one another (Ulmer et al. 2005) without forming any direct interhelical contacts (Bussell et al. 2005; Ulmer et al. 2005). Our data indicate that this topology is very likely to be preserved for both bS and gS.

Helix dynamics
Although the N-terminal domains of bS and gS are highly helical, as for aS, there is clear evidence that different regions of this helical structure are populated to different extents. Several regions in both proteins exhibit decreased indications of helical structure from chemical shifts, while at the same time showing increased intensities of both NOE crosspeaks and the associated diagonal signals, as well as a noticeable decrease in R2 and S2 values, indicating a greater degree of mobility. The combination of decreased helical content with increased motions most likely originates from fluctuations between highly helical and less helical conformations (Eliezer et al. 1998). This is seen most dramatically for bS in the second half of the second lipid-binding helix (past position 65), but is also evident for the N-terminal ~8 residues and between residues 30 and 40 for both proteins, as well as around position 65 and for positions 84–94 in gS. In this respect, gS closely resembles aS, which exhibits similar behavior for the N-terminal few residues, positions 30–40, around position 65 and at positions 84–94 (Bussell and Eliezer 2003; Bisaglia et al. 2005; Bussell et al. 2005; Ulmer et al. 2005). However, it is worth noting that positions 84–94 appear to be less dynamic in gS than in aS.

C-terminal tail
The C-terminal tails of both bS and gS are highly flexible and predominantly unstructured, as indicated by small and randomly fluctuating chemical shift deviations from random coil values, the absence of strong short range NOEs, and relaxation parameters consistent with an abundance of fast motions. Similar behavior was observed in the case of the C-terminal tail of aS (Bussell and Eliezer 2003; Chandra et al. 2003; Bussell et al. 2005), consistent with observations that there was little or no interaction between this tail and the surface of either detergent micelles or lipid vesicles (Eliezer et al. 2001). Nevertheless, there was some evidence of nonuniform behavior in the C-terminal tail of aS, with a small region around position 120 exhibiting aberrant C{alpha} chemical shift deviations, decreased dynamics, and an unusual response to the presence of paramagnetic Mn2+ ions. A similar region may exist around position 125 in bS, where a pronounced feature in the heteronuclear NOE data suggests a local decrease in flexibility, although there are no clear corresponding features in the chemical shift data. For gS, the C-terminal tail appears to be uniformly unstructured, with the flexibility increasing monotonically toward the very C terminus of the protein.


   Conclusions
TOP
 Abstract
 Introduction
 Results
 Discussion
 Conclusions
Materials and methods
 Electronic supplemental material
 References
 
Our results indicate that the major differences between the structures of aS, bS, and gS in the lipid-mimicking micelle environment correspond closely to differences in the primary sequences of the three family members. gS is more similar in sequence to aS in the repeat-containing lipid-binding domain, and our data confirm that the structural properties of the two proteins are very similar in this region, being highly helical with the previously observed break around position 42. However, gS differs structurally somewhat from aS in its acidic C-terminal tail, which is uniformly disordered, unlike that of aS, which appears to retain a region of less random structure. This difference reflects the lack of significant sequence conservation between the proteins in this region.

In contrast, the lipid-binding domain of bS differs notably from that of aS. While this domain remains predominantly helical in bS and preserves the break around position 42, there is a dramatic decrease in the stability of the helical structure between position 65 and position 83, where the helical region of bS ends. This change is clearly related to the deletion of 11 residues in the bS sequence, which serves to shorten the second helical stretch of the lipid-binding domain and appears to decrease its stability, resulting in greater dynamics and a lower population of the fully helical state. This effect might reflect a reduction in the lipid-binding affinity of this region of the protein. Although the deleted residues correspond to positions 74–84 in the aS sequence, the effects of the deletion appear to propagate as far as position 65 in the N-terminal direction and position 83 (corresponding to aS position 94) in the C-terminal direction. In both aS and gS, there is evidence of a structural perturbation in the helical structure around position 65 (Bussell and Eliezer 2003; Chandra et al. 2003; Bussell et al. 2005; Ulmer et al. 2005; present work). Combined with the data on bS, this suggests that the helical regions on either side of position 65 are somewhat decoupled from each other structurally. Thus, propagation of the perturbation of helical structure in bS is halted at this point. The C-terminal tail of bS is structurally more similar to that of aS than in the case of gS, reflecting the greater sequence conservation in this region between bS and aS. There is evidence for somewhat restricted motions in this region for both aS and bS, but not for gS.

Although the function of aS is not well understood, it is clear that at least one functional interaction of the protein, PLD2 inhibition, is mediated by the lipid-bound conformation of the protein (Payton et al. 2004). Differences in the micelle-bound conformations of the different synuclein family members could therefore determine how the functions of the three proteins differ. The structure of gS differs from that of aS primarily in the acidic C-terminal tail region. This region appears to play an important role in the protein–protein interactions of aS, and our results would therefore suggest that the interaction partners of aS and gS could differ significantly. In contrast, the primary structural differences between bS and aS occur in the lipid-binding domains of the two proteins. Therefore, it is likely that aS and bS will share many, though probably not all, protein-binding partners, and functional differences between the two proteins may be more strongly determined by their lipid interactions. Some support for these conclusions is provided by the fact that in the studies that originally identified PLD2 inhibition by synucleins, both aS and bS were identified as PLD2 inhibitors, whereas gS was not, despite the presence of all three proteins in the brain (Jenco et al. 1998).


   Materials and methods
TOP
 Abstract
 Introduction
 Results
 Discussion
 Conclusions
 Materials and methods
Electronic supplemental material
 References
 
Materials
Recombinant bS and gS were expressed in Escherichia coli BL21 (DE3) using plasmid constructs kindly provided by Dr. Peter Lansbury (Department of Neurology, Harvard Medical School). To produce isotopically labeled proteins for NMR studies, saturated overnight LB-kanamycin cultures were used to inoculate M9 minimal media made with uniformly labeled 13C-glucose and/or 15N-ammonium chloride. Cultures were grown at 37°C to an A600 nm of 0.5–0.6, at which point protein expression was induced with 1 mM IPTG. Cells were harvested by centrifugation 4 h post-induction. Proteins were purified using a protocol identical to that developed during previous studies of aS (Eliezer et al. 2001) involving ion-exchange and reverse-phase chromatography.

Circular dichroism
Circular dichroism (CD) spectra were measured on an AVIV 62 DS spectrometer equipped with a sample temperature controller. Far-UV CD spectra were monitored from 190 to 260 nm using final protein concentrations of 1 mg/mL with a path length of 0.2 mm, response time of 1 sec, and scan speed of 50 nm/min. Protein concentrations were measured using absorption at 280 nm. SDS micelle-bound protein samples were prepared under buffer conditions identical with those used in NMR experiments. Samples containing small unilamellar vesicles composed of a 1:1 ratio of 1-palmitoyl 2-oleoyl phosphatidylserine and phosphatidylcholine were prepared as previously described (Bussell and Eliezer 2004). Samples containing lysophosphatidylglycerol micelles (Krueger-Koplin et al. 2004) were prepared in a fashion identical to those containing SDS. Spectra were collected at 40°C.

NMR
NMR measurements were performed on samples dissolved in 100 mM NaCl, 10 mM Na2HPO4, 40 mM SDS (pH 7.4) in 90%/10% H2O/D2O as previously described for aS (Eliezer et al. 2001). NMR spectra were acquired at 40°C on a 600-MHz Varian Unity INOVA spectrometer. Pairs of HNCACB/CBCACONH and HNCACO/HNCO triple resonance experiments were collected for each protein to enable sequence-specific backbone and Cbeta resonance assignments. NOE measurements included NOESY-HSQC and HSQC-NOESY-HSQC experiments collected using 100 and 300 msec mixing times, respectively. To decrease spin diffusion effects, samples made with deuterated SDS were used in the NOESY experiments. 15N-edited HNHA spectra were collected to obtain 3JHNH{alpha} vicinal coupling constant values, with coupling constants calculated using the ratio of H{alpha} crosspeak intensities to diagonal resonance intensities (Vuister and Bax 1993). A dephasing/rephrasing delay of 25 msec was used and no correction for selective H{alpha} longitudinal relaxation rates was applied. Longitudinal (R1) and transverse (R2) relaxation rates for the backbone 15N nuclei were recorded using relaxation times of t1 = 10, 20, 40, 80, 160, 320, 640, 1280, and 1800 msec, and t2 = 14.4, 28.8, 43.2, 57.6, 115.2, 230.4, and 460.8 msec. To estimate noise levels, duplicate spectra were recorded for t = 80 and 640 msec (t1 spectra) and t = 14.4 and 115.2 msec (t2 spectra). R1 and R2 relaxation rates were determined by fitting resonance heights as a function of the relaxation delay time using NMRView (Johnson and Blevins 1994). 15N-(1H) steady-state heteronuclear NOE data were obtained as the ratio of peak heights in paired spectra collected with and without proton saturation during the relaxation delay of 5 sec. The experimental uncertainty was estimated using the standard deviation of four pairs of spectra. S2 values were derived from a model free analysis of the R1, R2, and heteronuclear NOE data using both the Modelfree (Mandel et al. 1995) and Tensor2 (Dosset et al. 2000) software packages, which gave similar results. Isotropic correlation times of 8.7 and 8.9 nsec were determined for bS and gS, respectively, using data from residues with a heteronuclear NOE value of 0.7 or greater (Tjandra et al. 1995). The difference between these values and that recently reported for aS (Ulmer et al. 2005) is caused by the different temperatures used (40°C vs. 25°C), which results in approximately a 60% reduction in 15N linewidths at the higher temperature for all three micelle-bound synuclein family members. The correlation times obtained were found to vary by approximately ±1 nsec, depending on the algorithm and heteronuclear NOE cutoff value used, but the effects of this variation on the S2 data were minimal and do not alter any of the results or conclusions. All NMR data were processed with NMRPipe (Delaglio et al. 1995) and analyzed using NMRView (Johnson and Blevins 1994). Resonance assignments have been deposited in the BioMagResBank database under deposition number 31871.


   Electronic supplemental material
TOP
 Abstract
 Introduction
 Results
 Discussion
 Conclusions
 Materials and methods
 Electronic supplemental material
References
 
Supplemental Figure S1 shows CD spectra of aS, bS, and gS bound to lysophospholipid micelles and phospholipid vesicles.


   Footnotes
 
Supplemental material: see www.proteinscience.org

Reprint requests to: David Eliezer, Department of Biochemistry and Program in Structural Biology, Weill Medical College of Cornell University, New York, NY 10021, USA; e-mail: dae2005@med.cornell.edu; fax: (212) 746-4843.

Article published online ahead of print. Article and publication date are at http://www.proteinscience.org/cgi/doi/10.1110/ps.051803606.


   Acknowledgments
 
This work was supported in part by the NIA, National Institutes of Health, grant AG19391 (to D.E.) and by a gift from Herbert and Ann Siegel (to D.E.). We thank Dr. Peter Lansbury for the kind gift of the bS and gS constructs and Trudy Ramlall and Carla Rospigliosi for technical assistance.


   References
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 Abstract
 Introduction
 Results
 Discussion
 Conclusions
 Materials and methods
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 References
 
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