The authors have declared that no competing interests exist.
Conceived and designed the experiments: AH JWG KM MEF SCK. Performed the experiments: JWG KM MEF SCK. Analyzed the data: JWG KM MEF SCK. Contributed reagents/materials/analysis tools: KN RBS. Wrote the paper: AH MEF RBS.
Synaptotagmin I (Syt I) is a vesicle-localized protein implicated in sensing the calcium influx that triggers fast synchronous release of neurotransmitter. How Syt I utilizes its two C2 domains to integrate signals and mediate neurotransmission has continued to be a controversial area of research, though prevalent hypotheses favor independent function. Using differential scanning calorimetry and fluorescence lifetime spectroscopy in a thermodynamic denaturation approach, we tested an alternative hypothesis in which both domains interact to cooperatively disseminate binding information. The free energy of stability was determined for C2A, C2B, and C2AB constructs by globally fitting both methods to a two-state model of unfolding. By comparing the additive free energies of C2A and C2B with C2AB, we identified a negative coupling interaction between the C2 domains of Syt I. This interaction not only provides a mechanistic means for propagating signals, but also a possible means for coordinating the molecular events of neurotransmission.
Regulated exocytosis of neurotransmitter requires the fusion of synaptic vesicles with the plasma membrane of the presynaptic neuron. This complex process is mediated by several key proteins including synaptobrevin, syntaxin-1, SNAP-25, complexins, and synaptotagmin I
How Syt I utilizes two C2 domains to rapidly transmute binding information from Ca2+ and lipid ligands as well as from proteins within its immediate vicinity to facilitate vesicle and plasma membrane fusion is not well understood, with conflicting evidence both for and against domain cooperation
Recent theoretical work
In light of the theoretical work above, we propose that rather than having an additive coincidence detector function, the C2 domains of Syt I cooperatively propagate binding information through inter-domain coupling. This type of interaction would be of significant functional relevance as each domain would become an allosteric regulator of the other. Ligand binding to C2B, for instance, would not only redistribute C2B’s ensemble, but would also increase or decrease the probability of binding-competent conformers of C2A being occupied.
Testing for the proposed interaction and quantifying its energetic value is difficult because of Δgint’s necessarily small magnitude. This problem is best appreciated in the context of the C2 domain’s restricted volume. The nine residue linker that tethers C2A and C2B together (
(A) Crystal structure of Syt I in the absence of ligand (PDB 2R83). The C2A domain, nine-residue linker region, and C2B domain are colored purple, blue, and green, respectively. (B) Conceptual representation and illustrative calculation of the volume accessible to the C2B domain with respect to C2A. Because the C2B domain is tethered to C2A, the volume it can occupy is restricted. This significantly increases the local concentration of C2B. If, for instance, the volume is calculated using the length of the linker and width of C2B as an approximate hemisphere radius (Pymol measurement of roughly 57 Å), the accessible volume is 4×10−19 cm3. Assuming 1 molecule of C2B occupies this volume, its local concentration is equal to (1 molecule C2B/6.022×1023 molecules per mole)/(4×10−25 m3) or ∼4 M. The local concentration of the C2A domain can be approximated in an analogous way. The resulting concentrations would shift the association equilibrium in the direction of bound domains.
Since our hypothesis is based upon subtle differences in protein energetics, two highly sensitive methods were employed to test it, namely differential scanning calorimetry (DSC)
To determine the relative stability of one domain compared to the other and whether or not the C2 domains of Syt I interact, the thermodynamic parameters of C2A and C2B in isolation were extracted from each domain’s thermal denaturation. The enthalpies measured for both C2A and C2B constructs did not show strong dependence on either concentration or scan rate consistent with the system reflecting equilibrium conditions (
C2B (Scan Rate, 1°C/min) | C2AB (Scan Rate, 1°C/min) | ||
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0.012 | 50.1 | 0.011 | 97.1 |
0.013 | 46.1 | 0.012 | 87.7 |
0.014 | 43.1 | 0.013 | 92.4 |
0.015 | 45.7 | 0.015 | 89.0 |
0.020 | 42.1 | 0.019 | 85.9 |
0.021 | 43.7 | 0.020 | 85.6 |
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45.1 | 2.9 | 89.6 | 4.4 |
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37.5–53.2 | 85.2–104.7 |
Footnote for
Circles represent the raw data and lines are the fitted model. C2A denaturations displayed with permission from Biophysical Journal. Higher concentrations refer to DSC samples. (A, E) [C2A] = 13 µM and 0.75 µM, [C2B] = 13 µM and 0.80 µM in the presence of 500 µM EGTA. (B, F) [C2A] = 13 µM and 0.75 µM, [C2B] = 12 µM and 0.75 µM in the presence of Ca2+. [Ca2+] = 800 µM and 770 µM for C2A, [Ca2+] = 5.3 mM and 5.2 mM for C2B. (C, G) [C2A] = 13 µM and 0.75 µM, [C2B] = 15.0 µM and 3.2 µM in the presence of LUVs consisting of 60∶40 POPC:POPS. [Lipid] = 870 µM and 50 µM for C2A, [Lipid] = 1.1 mM and 240 µM for C2B. Higher [C2B] in FLT was needed under lipid conditions to better see transition. (D, H) [C2A] = 13 µM and 0.75 µM, [C2B] = 13 µM and 3.2 µM for C2B in the presence of both LUVs and Ca2+. For C2A, [Ca2+] = 800 µM and [lipid] = 870 µM, [Ca2+] = 770 µM and [lipid] = 50 µM. For C2B, [Ca2+] = 5.2 mM and [lipid] = 290 µM, [Ca2+] = 5.2 mM and [lipid] = 70 µM. Low [lipid] for C2B prevented precipitation in calorimeter, but also limited FLT data collection due to flocculation and light scattering
C2B (Concentration 15 µM) | C2AB (Concentration 15 µM) | ||
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1.00 | 42.1 | 1.00 | 89.0 |
1.10 | 45.7 | 1.10 | 86.4 |
1.15 | 52.5 | 1.15 | 90.8 |
1.20 | 49.9 | 1.20 | 87.0 |
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47.5 | 4.6 | 88.3 | 2.0 |
Footnote for
Protein (Environment) | ΔHTm (kcal/mole) | ΔG° at 37°C (kcal/mole) | ΔS (kcal/mole·K) | Tm (°C) | ΔHTm/ΔHcal |
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58.7±0.3 | 2.32±0.05 | 0.18±0.01 | 56.0±0.1 | 0.98 |
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77.4±0.6 | 4.23±0.05 | 0.23±0.01 | 67.6±0.1 | 1.08 |
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61±1 | 2.44±0.05 | 0.19±0.02 | 55.5±0.2 | 1.03 |
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72.4±0.9 | 3.73±0.05 | 0.21±0.01 | 75.6±0.3 | 0.86 |
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69.6±0.6 | 1.74±0.09 | 0.22±0.01 | 46.4±0.1 | 1.45 |
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81.8±0.5 | 3.78±0.10 | 0.25±0.01 | 59.0±0.1 | 1.39 |
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63.7±0.1 | 1.13±0.02 | 0.20±0.01 | 43.3±0.1 | 1.51 |
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75.4±0.2 | 3.41±0.01 | 0.23±0.01 | 59.4±0.1 | 1.92 |
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102.9±0.4 | 2.24±0.01 | 0.32±0.01 | 45.6±0.1 | 1.05 |
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112.8±0.5 | 4.10±0.01 | 0.34±0.01 | 59.3±0.1 | 0.97 |
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86.8±0.4 | 1.74±0.01 | 0.27±0.01 | 45.1±0.1 | 1.05 |
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198±2 | 4.09±0.45 | 0.62±0.01 | 44.2±0.1 | NA |
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96.0±0.2 | 2.56±0.01 | 0.32±0.01 | 49.0±0.1 | 1.08 |
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136±1 | 5.53±0.02 | 0.41±0.01 | 58.0±0.1 | NA |
Footnote for
If C2A and C2B did not interact with one another when tethered together, the DSC denaturation profile would look similar to the superposition of both individual domain denaturations
(A) 10 µM of each domain in the presence of 500 µM EGTA. (B) 15 µM of each domain in the presence of 5.2 mM Ca2+ conditions. (C) 15 µM of each domain in the presence of 2.2 mM PS. (D) 11 µM of each domain in the presence of 2.9 mM PIP2. (E) 12 µM of each domain in the presence of 1.6 mM PS and 5.2 mM Ca2+. (F) 10 µM of each domain in the presence of 2.8 mM PIP2 and 5.2 mM Ca2+ conditions. Note that in most instances, two separate peaks are seen representing the independent unfolding of each domain. If C2A and C2B within the C2AB construct did not communicate, the C2AB denaturation profile would more closely resemble the above heat capacity profiles.
The Syt I C2AB construct was thermally denatured in the presence and absence of Ca2+ and lipid ligands. The enthalpy measured for this construct, like the individual domains, did not show a strong dependence on concentration or scan rate (
When comparing the additive free energies of the isolated C2A and C2B domains with the C2AB construct, a Δgint of −1.8±0.1 kcal/mol was found under ligand-free conditions at 37°C (Δgint = ΔG°C2AB – (ΔG°C2A + ΔG°C2B)). Since Δgint<0, the C2 domains of Syt I exhibit a negative inter-domain coupling interaction and the two domains experience opposite stabilizing effects upon introducing a perturbation. This form of coupling is similar to that of other allosteric proteins recently reported
The free energies of C2A, C2B, and C2AB in the presence of endogenous ligands were generally higher relative to a ligand-free environment (
The denaturation scans involving PIP2 differed between DSC and FLT methods (
Circles represent raw data and lines are the fitted model, excluding panels (H) and (I) wherein the line represents raw heat capacity data. Large and small concentrations refer to DSC and FLT concentrations, respectively. (A, D) 13 µM and 4.5 µM C2AB in the presence of 500 µM EGTA. (B, E) 12 µM and 0.75 µM C2AB in the presence of 5.2 mM and 5.1 mM Ca2+. (C, F) 12 µM and 0.75 µM C2AB in the presence of 1.7 mM and 110 µM PS. (G, J) 11 µM (3 replicates of DSC) and 0.75 µM C2AB in the presence of 2.9 mM and 210 µM PIP2. (H, K) 12 µM C2AB in the presence of 5.2 mM Ca2+ and 1.7 mM PS; 0.75 µM C2AB in the presence of 5.1 mM Ca2+ and 110 µM PS. (F, I) 11 µM (1 replicate of DSC) C2AB in the presence of 5.2 mM Ca2+ and 2.9 mM PIP2; 0.75 µM C2AB in the presence of 5.1 mM Ca2+ and 210 µM PIP2. Both calorimetric denaturations involving PIP2 had a limited number of replicates due to precipitation. In the absence of any interaction, the two domains would unfold independently. Instead, here the two domains unfold as one.
Solid, dashed, dotted, and dash-dot-dash lines represent EGTA, Ca2+, phosphatidylserine, and phosphatidylinositol environments. Note that most proteins of comparable size have maxima in the range of 10–20 kcal/mole
The purpose of the current study was to test a theory-driven hypothesis in which the C2 domains of Syt I interact to cooperatively disseminate binding information. In isolation, C2A and C2B were found to be energetically distinct. When tethered together, the two domains unfolded as one entity and were found to be less stable together than apart, indicative of a negative inter-domain coupling interaction. These results not only provide further experimental validation
In the absence of bound ligand (top), domains have basal level stability. When a ligand specific to the C2A domain binds (middle), C2A is stabilized and C2B is destabilized through negative coupling. When a ligand specific to C2B binds (bottom), the opposite effect is seen; C2B is stabilized and C2A is destabilized. Note that binding of domain-specific ligands lowers the probability of binding-competent conformers being occupied in the adjacent domain through domain destabilization, representing a form of allosteric regulation. The extent of negative coupling, like domain stability, changes depending on the types of ligand present.
Aside from showing negative inter-domain coupling, the results presented here also indicate that the interaction is subject to modulation by Ca2+ and phospholipid ligands. The general observation of marginal stability in all three protein constructs relative to proteins of similar size indicates that Syt I has a larger degree of conformational freedom and thus is more flexible. Additionally, the isolated C2A and C2B domain free energies of stability (which report on the overall breadth of each domain’s conformational ensemble) in the presence and absence of various ligands indicates a high degree of malleability (
The differential binding preferences of each C2 domain can, however, modulate Δgint. In the presence of PS alone (which had an overall destabilizing effect, decreasing C2AB’s free energy from 2.24 kcal/mole to 1.74 kcal/mole), Δgint did not change within error (went from −1.8 kcal/mole to −1.83 kcal/mole). Here, the binding of PS to C2A does not appear to have as great of a destabilizing effect on the adjacent C2B domain. If, however, Ca2+ ligand is added to drive the C2A domain into a lipid bound state, the degree of C2B destabilization becomes more pronounced. Evidence for this is seen in the FLT denaturations of C2AB under PS and Ca2+ conditions (
Two of three tryptophan residues (C2A’s and one of C2B’s) occupy superficial positions and may, as a result, be more solvent exposed in solution (left). The second tryptophan in the C2B domain is partially embedded in the core of the β-sandwich motif amongst several hydrophobic residues (yellow) (right). The differences in tryptophan environment likely give rise to unequal FLT signal contributions, with most of the signal coming from C2B’s β-sandwich residue.
DSC denaturations of C2AB under the analogous PS and Ca2+ conditions lend further support to this negative coupling hypothesis. C2B, presumably the first peak in the heat capacity profile of
In contrast to the change in free energy brought about by binding of PS, PIP2 had an overall stabilizing effect in the C2AB construct (as indicated by the increase in free energy from 2.24 to 2.56 kcal/mol). There were, however, discrepancies between DSC and FLT denaturation profiles. We attribute these differences to the sensitivity of FLT to microenvironment. Because PIP2 is a specific ligand for C2B, its binding might select for a subset of conformers in which more water is excluded from the β-sandwich interior. This in turn might make the tryptophan residue that resides there (Trp 390) more sensitive to changes in solvation in response to increasing temperature and thus account for an early unfolding transition relative to DSC. In the case of PIP2 and Ca2+ where the DSC profile is somewhat suggestive of independent domain unfolding, not seeing two independent transitions on FLT may be the result of unequal tryptophan signal contribution. Because PIP2 is a ligand for C2B, it binding would presumably destabilize C2A through negative coupling, an effect perhaps accentuated by the additional presence of calcium ion
In this denaturation study, two distinctly different techniques were used to monitor the unfolding transition of each protein construct. The intent of using both DSC (which provides a global perspective) and FLT (a more local perspective) was to overcome any inherent bias of using a single technique in isolation. When comparing the global fit of all DSC and FLT replicates to each individual method, minor deviations of the model were observed in DSC plots of the individual C2 domains. These observations suggest that, indeed, neither technique fully captures the unfolding transition on its own. In the case of C2AB, the unique perspective of each denaturation method is particularly prominent, as noted above. By including both DSC and FLT data sets in the global fit, these unfolding transitions are more completely described. Furthermore, the global fit approach is validated by the close agreement of calculated and calorimetric enthalpies (ΔHTm/ΔHcal ratio in
When considering the work presented here, it is important to note how the approach used and the results obtained differ from those of previous experimentation looking at domain interaction. Both the free energy of stability and free energy of interaction are global perspectives of Syt I behavior. As such, any specific structural contact points
With these distinctions in mind, the above observations and thermodynamic profiles can introduce additional insight into Syt I function. The small energetic values of both ΔG°37°C and Δgint suggest sensitivity not only to endogenous ligands, but also structural changes within the protein, like those that arise from gene mutation. Indeed, Syt I’s functional sensitivity to mutation has been well documented
In the context of normal neurotransmission, the large and malleable conformer ensemble of Syt I has further functional implications. If the conformer ensembles of C2A and C2B are subject to modulation by ligands, they may, by extension, be influenced by other domain-specific binding partners in the immediate vicinity within the cell. Indeed, recent EPR and FRET studies looking at Syt I interactions with SNARE proteins show that even when bound to the SNARE complex, structural fluctuation and conformer heterogeneity still exist, indicating yet another possible means for modulating the Syt I conformer ensemble
Potassium chloride (KCl) was Puriss-grade. Calcium chloride dihydrate, 3-(N-morpholino) propanesulfonic acid (MOPS), and ethylene glycol-bis(2-aminoethyl)-N,N,N′,N′tetra-acetic acid (EGTA) were all Biochemika grade from Fluka Chemical Corp. All buffers were decalcified using Chelex-100 ion-exchange resin (Bio-Rad Labs). All lipids including 1-palmitoyl-2-oleoyl-
Large unilamellar vesicles (LUVs) consisting of POPC:POPS (60∶40) and POPC:PIP2 (95∶5) were prepared as previously described
Human Syt I C2A and C2AB constructs were purified according to previously described methods
DSC experiments were performed on a NanoDSC (TA Instruments, New Castle, DE) at a scan rate of 1°C/min. To see if measured enthalpies varied with concentration or scan rate, all constructs were denatured over a range of concentrations and scan rates
FLT experiments were performed on a Lifetime Spectrometer (Fluorescence Innovations, Bozeman, MT) using nanomolar protein concentrations. Under some ligand conditions, higher concentrations were needed to obtain good signal. Scans were conducted in chelexed 20 mM MOPS, 100 mM KCl, pH 7.5. No time-resolved measurements were made. Instead, the integrated intensity of the lifetime decay was used to selectively monitor intrinsic fluorescence of endogenous tryptophan residues (excitation and emission wavelengths of 295 and 340 nm, respectively). Change in fluorescence emission for each construct was monitored as a function of increasing 2°C temperature increments. In scans free of Ca2+, 500 µM EGTA was added as background. In scans with Ca2+, both C2A and C2B binding sites were >95% saturated (as described above in DSC section). For C2AB, a Ca2+ concentration of 5.1 mM was used. The same Ca2+ stock solution described above was used for FLT samples. Scans with lipid contained LUVs of identical composition as described above for DSC. Percent reversibility was measured by comparing the integrated fluorescence lifetime intensity of the sample before heating and after cooling. All data sets were analyzed at an emission wavelength of 345 nm to verify absence of contributing water fluorescence at 340 nm. The fluorescence signal measured was normalized to the calculated intensities from the two-state model and subsequently displayed as “Normalized Intensity” in each corresponding plot.
The free energy of stability for each construct under each set of ligand conditions was determined by globally fitting denaturation data from DSC and FLT methods to a two-state transition model. For full details of this model and its application, see
The equilibrium constant can be used to represent fractions of folded (fN) and unfolded (fD) protein throughout the transition. As a protein undergoes thermal denaturation in the DSC, the heat capacity (Cp(T)) of the sample cell changes as the fraction of unfolded protein changes:
Where ΔH(T) is the associated enthalpy. When the protein is denatured in FLT, the tryptophan residues become more solvent exposed and lose much of their initial intensity. Throughout the unfolding transition, the total fluorescence signal measured (S(T)) is a composite of native (SN) and denatured (SD) protein fluorescence and depends on the fraction of each present at a given temperature:
Where SN and SD are approximated by using linear equations. By substituting the Gibbs-Helmholtz equation (
The terms ΔHTm, Tm, and ΔCp are the fit parameters in this model, however, to further constrain this fit, ΔCp was fixed using an empirical approximation method that showed good agreement with experimentally-derived values
Lastly, the entropy change associated with the unfolding transition (ΔS) was calculated using the ΔG°37°C, ΔHTm, and the physiological temperature of 310 K of each protein construct under each set of ligand conditions using the Gibbs free energy equation:
Application of the fluctuation dissipation theorem to C2AB. Qualitative representation of native (N) and denatured (D) enthalpy distributions for C2AB in the absence of any ligand (blue), in the presence of POPC:POPS (60∶40) liposomes (green), and in the presence of Ca2+ (red). Note that the enthalpy distributions of an N/D pair shift in response to ligand, consistent with C2AB malleability and a ligand-induced change in conformer distribution.
(TIF)
Effectiveness of the C2B purification protocol used for this study. Prominent band in lanes 2, 3, and 5 are MBP-C2B after cell lysis (2), during elution of unwanted protein (3), and during 250 mM imidazole elution (5). Lane 6 shows separation of MBP from C2B. Lane 7 shows pure C2B after passing cut MBP-C2B over column. Ladder (lanes 1 and 8, Precision Plus, Bio-Rad Labs) has molecular weights (kDa) of 10, 15, 20, 25, 37, 50, 75, 100, 150, and 250.
(TIF)
Complete list of calorimetric enthalpies used to assess concentration dependence of the C2B domain.
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Complete list of calorimetric enthalpies used to assess concentration dependence of the C2AB cytosolic fragment.
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Thermodynamic parameters and associated errors for C2B and C2AB using FLT λem of 345 nm. Analogous controls were performed for C2A. When FLT integrated fluorescence intensity was normalized and globally fit, there was no substantial variation in calculated fit parameters. This indicated that water fluorescence (water raman at 328 nm) at 340 nm was negligible.
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Two-state derivation, heat capacity fluctuation dissipation theorem, C2B purification, and non-linear least squares regression analysis.
(DOC)