Insight into the Conformational Transitions of Serine Acetyl Transferase Isoforms in E. histolytica: Implications for Structural and Functional Balance.

Serine acetyl transferase (SAT) is one of the crucial enzymes in the cysteine biosynthetic pathway and an essential enzyme for the survival of Entamoeba histolytica, the causative agent of amoebiasis. E. histolytica expresses three isoforms of SAT, where SAT1 and SAT2 are inhibited by the final product cysteine, while SAT3 is not inhibited. SAT3 has a slightly elongated C-terminus compared to SAT1. To understand the stability and conformational transition between two secondary structures of proteins, we measured the effect of urea, a chemical denaturant, on two isoforms of SAT (SAT1 and SAT3) of E. histolytica. The effect of urea on the structure and stability of SAT1 and SAT3 was determined by measuring changes in their far-UV circular dichroism (CD), Trp fluorescence, and near-UV absorption spectra. The urea-induced normal transition curves suggested that the structural transition is reversible and follows a two-state process. Analysis of the urea-induced transition of all optical properties for the stability parameters ΔG D° (Gibbs free energy change (ΔG D) in the absence of urea), m (dependence of ΔG D on urea concentration), and C m (midpoint of urea transition) suggested that SAT1 is more stable than SAT3. Characterization of the end product of the urea-induced transition of both proteins by the far-UV CD and Trp-fluorescence and near-UV absorbance suggested that urea causes α-helix to β-sheet transition and burial of Trp residues, respectively. To support the in vitro findings, 100 ns molecular dynamics simulations (in silico study) were performed. Both the spectroscopic and molecular dynamics approaches clearly indicated that SAT1 is more stable than SAT3. SAT3 has evolved to escape the feedback inhibition to keep producing cysteine, but in the process, it compromises its structural stability relative to SAT1.

Introduction

Entamoeba histolytica is an enteric protist parasite that causes amebic colitis and extra intestinal abscesses (i.e., hepatic, pulmonary, and cerebral).[1,2] The cysteine biosynthetic pathway is important for the growth, survival, and pathogenicity of E. histolytica as it is involved in the synthesis of cysteine which serves crucial roles as antioxidative agents and as a sulfur source for various biomolecules like thiols (glutathione, trypanothione, mycothiol), biotin, Fe–S clusters, and thiamine.[1,3] Serine acetyltransferase (SAT1 EC 2.3.1.30) is an enzyme of the l-cysteine biosynthesis pathway, which catalyzes the acetyl-CoA dependent acetylation of the side chain hydroxyl group of l-serine to form O-acetylserine. Another enzyme of the pathway is O-acetyl serine sulfhydrylase (OASS) that further catalazes the insertion of sulfide into OAS and generate cysteine with the release of acetate.[4] The cysteine biosynthetic pathway is regulated by several mechanisms, including feedback inhibition of SAT by cysteine.[1,5]E. histolytica has three isoforms of SAT (SAT1, SAT2, and SAT3). SAT1 shows 78% sequence homology with SAT2 and 48% with SAT3. These three isoforms are differing in length and show differences in sensitivity to the feedback inhibitor l-cysteine.[4] SAT1 and SAT2 are inhibited by Cys via a feedback mechanism, although SAT3 is insensitive to this inhibition.[6]

Biological studies indicate that SAT3 is very crucial for survival, while SAT1 and SAT2 deleted mutant strains can still survive.[7] SAT1 is a trimeric protein, and its protomer structure contains two domains, an N-terminal α-helix-rich domain composed of a collective of eight α-helices responsible for oligomerization and a C-terminal left handed β-helical domain comprised predominantly of 14 β strands.[8] Like SAT1, SAT3 is also trimeric, and its N-terminal domain is highly variable in structure and charge distribution, which plays a role in not forming hexameric structures as seen in E. coli and other structures. The C-terminal domain is highly conserved in all organisms and plays a role in trimerization,[8] while the C-terminal end is highly diverse. Especially E. histolytica SAT3 has a long C-terminal end which is swapping over other monomers.[8]

Most of the folding studies have been carried out on the monomeric proteins as a model to understand the folding pathway of proteins in vitro.[9,10] However, complicated oligomeric F proteins are also used to study the folding pathway.[11,12] The major secondary structural elements are β-sheets and α-helices that facilitate the organization of the three-dimensional (3D) conformation of proteins and peptides.[13] The proper folding of the protein is to adopt the native structure to perform normal functions, whereas misfolding is linked to malfunctions and aggregation as well as various neurodegenerative diseases in humans.[14,15] The conformational changes in human protein structure from α-helix to β-sheets are responsible for various neurodegenerative diseases.[16−18] However, understanding of the structural behavior of β-sheet assemblies in neurodegenerative diseases remains poorly understood.

Despite being vital for the survival of E. histoyltica, very little information is available on the stability and biophysical properties of SAT isoforms. In this study, we report biophysical and thermodynamic properties of SAT1 and SAT3 isoforms of E. histolytica measured by different spectroscopic techniques that were used to monitor the urea-induced transition of these isoforms. The structural characterization of the end product of this chemical denaturant-induced transition suggested that each isoform undergoes an α-helical → β-structure transition resembling neurodegenerative structural transition. 100 ns MD simulations at 300 K were also carried out in water and different concentrations of urea to establish a correlation between in vitro and in silico results. Both the spectroscopic and molecular dynamics studies clearly indicated that SAT1 is more stable than SAT3.

Materials and Methods

Materials

Ultrapure-grade urea was purchased from MP Biomedicals (India) Pvt. Ltd. Oligonucleotide primers were supplied by Polaris Biosciences India Pvt. Ltd. Agarose, bacto-tryptone, and yeast extract were purchased from HiMedia Laboratories, India. E. coli strain DH5α was used for DNA amplification, and the Escherichia coli BL21 (DE3) strain was used for expression of the proteins. pET-21c, and pET-28b (Novagen, Wisconsin, USA) were used as expression vectors. DNA restriction enzymes (Nhe1 and Xho1) and T4 DNA-ligase were purchased from Thermo Scientific, USA. Phusion and Dnazyme DNA polymerase (Taq polymerase) were obtained from Promega. Lysozyme was obtained from Amersham Pharmacia Biotech. All chemicals and reagents were of analytical grade and highly pure and were, therefore, used without further purifications.

Expression and Purifications

SAT1 and SAT3 genes were subcloned in pET-21c and pET-28b expression vectors, respectively. These constructs were transformed into Escherichia coli BL21 (DE3) cells, and freshly transformed Escherichia coli BL21 (DE3) cells were grown in LB media supplemented with 100 μg/mL ampicillin at 37 °C to an A600 (absorbance at 600 nm) of 0.6. Then cultures of both SAT1 and SAT3 were induced by 0.5 mM isopropyl β-d-1-thiogalactopyranoside (IPTG) for overexpression of proteins, and cultures were allowed to grow for another 3 h at 37 °C. Cells were harvested by centrifugation at 7000g for 7 min at 4 °C. The harvested cells were suspended in lysis buffer (50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 5% (v/v) glycerol, 5 mM β-mercaptoethanol, 0.1 mg/mL lysozyme, 1 mM phenylmethanesulfonyl fluoride containing 1% (v/v) Triton X-100 and lysed with three cycles of flash-freezing in liquid nitrogen and thawing. The cell lysate was sonicated on ice and centrifuged at 12 000g for 30 min at 4 °C. The supernatant was collected and subjected to Ni-NTA affinity chromatography. Further, proteins were purified by gel filtration chromatography.[4,8]

Preparation of Stocks Solutions of Urea and Sample Preparation

The stock solution of 10 M urea was prepared in 50 mM Tris buffer at pH 8.0. The urea stock solution was always freshly prepared to avoid the production of cyanate ions, and its concentration was confirmed by refractive index.[19] For chemical-induced transition studies of SAT1 and SAT3, protein solutions with different concentrations of urea were prepared in a volumetric flask and incubated overnight, enough time for the completion of the transition.[20] A concentration of 8.6 μM of SAT1 and 7.90 μM of SAT3 proteins was used in all measurements. Urea was added in subsequent samples with a gradual increase in concentration from 0.25 to 8.0 M. The urea-induced transition was studied at pH 8.0 and 25 °C.

Circular Dichroism (CD) Measurements

Secondary structures of SAT1 and SAT3 were measured with a Jasco CD spectrometer (model J-1500) equipped with a Peltier type temperature controller. The far-UV CD spectra (250–200 nm) were recorded at 25 ± 0.1 °C using a cuvette of 1 mm path length. The protein concentration of both proteins used in these measurements was 8.6 μM of SAT1 and 7.90 μM of SAT3 in a buffer of 50 mM Tris at pH 8.0 and 150 mM NaCl. To monitor the effect of urea on the secondary structure of SAT1 and SAT3, far-UV CD spectra of each protein were recorded in the presence of different concentrations of the denaturant.[21] The recorded CD spectra were uploaded onto the K2D2 server to estimate the secondary structural content of the protein.[22] This server is based on the comparison of real and predicted values, by means of the Pearson correlation coefficient (r) and the root-mean-square deviation (RMSD) of proteins with known 3D structure. The raw CD data were converted to the mean residue ellipticity, [θ] (degree cm2 dmol–1), using the relation:where θλ is the observed ellipticity in millidegrees at wavelength λ nm, M0 is the mean residue weight of the protein, c is the concentration of the protein in mg/mL, and l is the path length of the cell in cm.

Fluorescence Measurements

Fluorescence spectral measurements were carried out in a Jasco spectrofluorimeter (Model FP-6200) using a 3 mm quartz cell at 25 ± 0.1 °C. These measurements provide information on the tertiary structures of SAT1 and SAT3. Both excitation and emission slits were set at 5 nm bandwidth. The temperature of the sample cell was maintained by circulating water using an external thermostated water bath. In these experiments, protein samples were excited at 292 nm, and emission spectra were recorded in the range of 300–400 nm. The protein concentration used was 8.6 μM of SAT1 and 7.90 μM of SAT3, respectively. To see the effect of urea on the tertiary structure of SAT1 and SAT3, intrinsic (Trp) fluorescence spectra were recorded as a function of the denaturant concentration.

Absorption Measurements

Near-UV absorption spectra of SAT1 and SAT3 were measured in a Jasco UV/visible spectrophotometer (Jasco V-660, Model B028661152) equipped with Peltier temperature controller (ETCS-761). To monitor the effect of urea on the tertiary structures of SAT1 and SAT3, all spectra were recorded in the wavelength range 340–240 nm. For each measurement, 8.6 μM of SAT1 and 7.90 μM of SAT3 concentration of proteins were used. Difference spectra were obtained by subtracting the spectrum of the native protein from the spectrum of the protein in the presence of denaturants. Absorbance at a given wavelength λ nm was converted into the molar absorption coefficient (ελ). A transition curve was generated by plotting Δε287 as a function of [urea], the molar concentration of urea. For each sample, at least three independent measurements were performed and averaged for analysis.

Data Analysis

The isothermal transition curves of SAT1 and SAT3 induced by urea were analyzed assuming that the transition process is reversible and proceeds through a two-state mechanism (i.e., N (native) state ↔ D (denatured) state). [θ]222 was used to monitor the change in secondary structure, whereas Δε287 and F345 (fluorescence intensity at 345 nm) indicate the changes in the tertiary structure of the protein as an influence of urea.[19] All three optical properties ([θ]222, Δε287, and F345) were plotted against [urea] to obtain the transition curves. Each transition curve was analyzed for the estimation of stability parameters[23] using a nonlinear least-squares method according to eq .[24] Transition curves of all three optical methods were fitted according to eq and give the values of C and m.where y(d) is the optical property observed at [d], the molar concentration of the denaturant; yN(d) and yD(d) are optical properties of the native and denatured protein molecules at [d], respectively; ΔGD0 is the value of the Gibbs free energy change (ΔGD) in the absence of the denaturant; md is the slope (∂ΔGD/∂[d]); R is the universal gas constant; and T denotes the temperature in Kelvin. It should be noted that the analysis transition curve was done assuming that unfolding is a two-state process and reversible and dependencies of yN(d) and yD(d) are linear (i.e., yN(d) = aN + bN[d] and yD(d) = aD + bD[d], where a and b are [d]-independent parameters, and subscripts N and D represent that these parameters are for the native and denatured protein molecules, respectively. At a given denaturant concentration, the fraction of the protein converted to β-structure fβ was estimated using the relation:

Molecular Dynamics Simulations

Molecular dynamics (MD) simulations of both SAT1 and SAT3 (three-dimensional structure was retrieved using the PDB ID 3PIB and PDB ID 7BW9, respectively) were performed using the GROMACS 2018.4 package. Both proteins were simulated in the presence of 4.0, 6.0, and 8.0 M urea. Before MD simulations, the structures were converted to GROMACS supported formats using the PDB 2gmx tool of GROMACS. The topology files were generated using the same tool with the force field GROMOS96 43a1. The structures then solvated using the aforementioned concentrations of urea, and water molecules were added. Subsequently, Na+ and Cl– ions were added in the system. Both the systems were then energy minimized using the steepest descent algorithm implemented in GROMACS for 100 ps (ps). Both the structures were analyzed for proper energy minimization. The energy minimized systems were then taken to equilibration using NVT and NPT methods for 100 ps each. The equilibrated systems were then considered for production MD of 100 ns (ns). Eventually, SAT1 in aqueous solution and in the presence of 4.0, 6.0, and 8.0 M urea were simulated for 100 ns. Similarly, SAT3 under the same solvent conditions was simulated for 100 ns. The trajectories generated after the 100 ns MD simulation of each system were then analyzed for structural changes and stability.

Results and Discussion

Protein structure is crucial to its biological function. The various molecular interactions maintain the protein’s proper folding and stability under physiological conditions. It is observed that cells and tissues under stress conditions due to a change in the solute concentration, high temperature, and some of the metabolic substances affect the activity and stability of proteins. The common chemical denaturants such as urea and GdmCl are used to induce the unfolding of protein to estimate the protein stability parameters. The protein folding and stability of monomeric protein studies have been carried out extensively. But, in recent years, the folding and stability of oligomeric proteins have been carried out to understand the structure–function relationship. In vitro and MD simulation approaches have been used to understand the mechanism of transition of proteins by urea.[20,25] In the present study, effects of urea on the stability of SAT1 and SAT3 were monitored using three optical methods and an MD simulation approach, which correlated very well. These results clearly indicate that SAT1 is more stable than SAT3, while functionally SAT3 is important for the survival of the organism.

Expression and Purifications of SAT1 and SAT3

SAT1 and SAT3 were expressed in Escherichia coli BL21 (DE3) cells, and these cells were harvested and resuspended in a lysis buffer. The cell lysate was sonicated on ice and centrifuged at 12 000g for 30 min at 4 °C. The supernatant was collected and subjected to Ni-NTA affinity chromatography. A clear supernatant of each SAT1 and SAT3 was passed through a Ni-NTA column, which was pre-equilibrated with the equlibration buffer (50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 5% (v/v) glycerol, 5 mM β-mercaptoethanol, and 10 mM imidazole). This column was washed with five volumes of the wash buffer with 20 mM imidazole. The bound protein was eluted with 15 mL of elution buffer with 300 mM imidazole and collected in 1.5 mL fractions. The purity of eluted fractions of both proteins was checked on SDS-PAGE, and some impurities were found along with SAT1 and SAT3 proteins. Furthermore, proteins were subjected to gel filtration chromatography.[20,26] The proteins were concentrated using a centricon filter with a 30 kDa cutoff and loaded onto a HiLoad 16/60 Superdex 200 column (GE Healthcare) that had previously been equilibrated with a buffer of 50 mM Tris at pH 8.0, 150 mM NaCl, and 5% glycerol. Proteins were eluted at a flow rate of 1 mL/min. SAT1 was eluted at 72 mL, while SAT3 was at 84 mL. The elution profile shows that SAT1 and SAT3 exist as a trimer (Supplementary Figures S1 and S2). The purity of the proteins was checked on SDS-PAGE and concentrated using Centricon tubes (Amicon) to a final concentration of 12 mg/mL for SAT1 and 8 mg/mL for SAT3 as estimated by the Bradford method.

CD Measurements

The far-UV CD measurement is used to determine secondary structure.[27,28] It is also used to follow the transition between α-helix and β-structure. In the presence of different concentrations of urea, far-UV CD measurements were performed on SAT1 and SAT3 at 25 °C in the wavelength region 250–200 nm (Figure A and C). It should be noted that measurement of the CD spectrum of proteins is not possible beyond 215 nm due to a high photomultiplier voltage. Figure A and C show representative spectra, and other spectra are shown in Figure S2A and B (Supporting Information). The CD spectrum of each native protein was analyzed for the content of α-helix, β-structure, and unordered structure (random coil) using an online available program K2D2.[22] The analysis of the CD spectrum of the native SAT1 gave values of 20.2% for α-helix, 29.1% for β-sheet, and 50.7% random coil.[29] Analysis of the CD spectrum of the native SAT3 yielded values of 22.2% for α-helix, 27.1% for β-sheet, and 50.3% for random coil. Values of the secondary structural elements are in close agreement with those observed in the crystal structure.[26] These agreements mean that our CD measurements are accurate. We did not attempt to analyze CD spectra of proteins in the presence of the denaturant, for these spectra cannot be accurately obtained beyond 215 nm. It is seen in Figure A and C that, on the addition of urea, SAT1 and SAT3 undergo α → β transition.

Figure 1

Far-UV CD spectra of SAT1 and SAT3 recorded as a function of increasing urea concentration at pH 8.0 and 25 °C. (A) Representative far-UV CD spectra of SAT1 in the presence of different concentrations of urea. (B) Urea-induced transition curve of SAT1 followed by observing changes in [θ]222. (C) Representative far-UV CD spectra of SAT3 in the presence of different concentrations of urea. (D) Urea-induced transition curve of SAT3 followed by observing changes in [θ]222.

Values of [θ] at 222 nm were obtained from CD spectra shown in Figure A and C and plotted against [urea]. These plots (urea-induced transition curves) for SAT1 and SAT3 are shown in Figure B and D, respectively. It is seen in these figures that SAT3 transition starts above 2.0 M urea and is complete around 5.5 M urea (Figure D), and, on the other hand, SAT1 undergoes transition from 2.0 M urea, which is complete above 6.5 M urea (Figure B). Thus, it seems that SAT1 is more stable than SAT3. As may be seen in Figure B and D, there is no change in the secondary structure of SAT1 and SAT3 up to ∼2.5 M urea concentration, but the loss of secondary structure gets started above this concentration, and it continues up to 5.25 M urea. After that, no further change was observed for these proteins. The transition curves of SAT1 and SAT3 show that with an increase in urea concentration, a cooperative transition resulted. We have analyzed the transition curves of both proteins for ΔGDo, m, and Cm according to eq . The thermodynamic parameters are given in Table .

Table 1

Thermodynamic Parameters Obtained from Urea-Induced Denaturation of SAT1 and SAT3 at pH 8.0 and 25 ± 0.1 °C

protein	SAT1			SAT3
probes (urea)	ΔGD°	m	Cm	ΔGD°	m	Cm
[θ]222	5.91 ± 0.17	2.50 ± 0.11	2.35 ± 0.15	3.93 ± 0.13	1.09 ± 0.08	3.61 ± 0.16
F345	4.85 ± 0.13	1.25 ± 0.09	3.88 ± 0.14	4.01 ± 0.15	1.02 ± 0.07	3.93 ± 0.20
Δε287	5.13 ± 0.15	1.37 ± 0.07	3.74 ± 0.17	3.88 ± 0.14	1.11 ± 0.04	3.40 ± 0.19
average	5.29 ± 0.15	1.70 ± 0.09	3.32 ± 0.15	3.94 ± 0.14	1.01 ± 0.06	3.65 ± 0.18

Tryptophan fluorescence spectral studies provide information on the tertiary structure of a protein. The fluorescence spectra of SAT1 and SAT3 were measured in the presence of different concentrations of urea at pH 8.0 (50 mM Tris buffer) and 25 °C. The hydrophobic core of globular or multidomain proteins is folded in such a way that tryptophan residues are preferably buried and protected from the solvent. Figure A and C show representative fluorescence spectra, and other spectra are shown in Figure S3A and B (Supporting Information). It is seen in these figures that the intensity of the emission peak of SAT1 and SAT3 increased with a blue shift with an increase in the urea concentration. A blue shift of approximately 5 nm was observed. There are two tryptophan residues in SAT1 (Trp195 and Trp255) and three in SAT3 (Trp195, Trp255, and Trp308). Trp195 and Trp255 are buried in both proteins, while Trp308, which is not present in SAT1, is exposed. The observed increase in fluorescence intensity with a blue shift in urea solutions suggests that Trp residue(s) is (are) transferred to a more nonpolar environment.

Figure 2

(A) Representative fluorescence spectra of SAT1 recorded as a function of increasing urea concentration at pH 8.0 and 25 °C. (B) Urea-induced transition curve followed by observing changes in F345 of SAT1. (C) Representative fluorescence spectra of SAT3 recorded as a function of increasing urea concentration at pH 8.0 and 25 °C. (D) Urea-induced transition curve followed by observing changes in F345 in SAT3.

Figure B and D respectively show urea-induced transition curves of SAT1 and SAT3 monitored by F345 (Trp-fluorescence emission intensity at 345 nm). The transition curves were analyzed to estimate the values of ΔGDo, m, and Cm according to eq . These thermodynamic parameters are listed in Table .

Near-UV absorbance spectral measurements provide information on the tertiary structure of a protein. Absorption spectra of the SAT1 and SAT3 with different concentrations of urea were measured at pH 8.0 (50 mM Tris buffer) and 25 °C to study the effect of this chemical denaturant on the tertiary structure of both proteins. Figure A and C shows representative absorption spectra, and other spectra are shown in Figure S4A and B (Supporting Information). It is seen in these figures that the addition of urea to the protein solution causes an increase in the absorbance near 280 nm. It seems that, as observed from the fluorescence measurements (Figure A and C), Trp residue(s) is (are) transferred to a more nonpolar environment on treating proteins with urea.

Figure 3

Near-UV absorption spectra of SAT1 and SAT3 at pH 8.0 and 25 °C. (A) Representative absorption spectra of SAT1 recorded as a function of increasing urea concentration. (B) Transition curve of SAT1 obtained by plotting the Δε287 value as a function of [urea]. (C) Representative absorption spectra of SAT3 recorded as a function of increasing urea concentration. (D) Transition curve of SAT3 obtained by plotting the Δε287 value as a function of [urea].

Figure B and D respectively show urea-induced transition curves of SAT1 and SAT3, monitored by Δε287 (difference in molar absorption coefficient in the presence and absence of urea at 287 nm). These transition curves (plots of Δε287 versus [urea]) were analyzed for ΔGDo, m, and Cm according to eq . These thermodynamic parameters are listed in Table .

The isothermal transition of SAT1 and SAT3 induced by chemical denaturant urea were carried out to estimate the protein stability. Normalized transition curves of different physical properties of proteins are used as a test for the two-state transition. The transition curves shown in Figures B and D, 2B and D, and 3B and D were normalized to estimate fβ at different urea concentrations using eq . These values of fβ are plotted against the concentration of urea in Figure (A and B) that shows a coincidence of normalized transition curves.

Figure 4

Normalized urea-induced transition curves of SAT1 and SAT3 at pH 8.0 and 25 °C. (A) SAT1 and (B) SAT3. Data were obtained from the analysis of transition curves of Δε287, F345, and [θ]222 to construct plots of fβ of SAT1 and SAT3 versus [urea].

In the analysis of urea-induced transition, a few assumptions were made. The first assumption is that the urea-induced transition follows a two-state mechanism. The normalized curves of three optical properties for SAT1 and SAT3 against [urea] (Figures , 2, and 3) coincide with each other as shown in Figure A and B, respectively suggesting that the first assumption is correct. Furthermore, analysis of transition curves shown in Figures –3 according to eq gave identical values of ΔGDo, m, and Cm for a protein (see Table ). Almost similar values of each thermodynamic parameter obtained from the analysis of transition curves of three different optical properties support the hypothesis that urea-induced transition of SAT1 and SAT3 follows a two-state mechanism.

The second assumption is that ΔGD-dependence on [urea] is linear in the entire range. This assumption seems to be valid for many proteins.[30,31] The third assumption is that the optical properties of the protein in the native state (yN) are independent of [urea], and the denatured state (yD) is dependent on [urea] in a linear manner.

These stability measurements of SAT1 and SAT3 indicate that SAT1 is more stable than SAT3. CD measurements suggested that urea induces α → β transition in both proteins. Trp fluorescence and near-UV absorption measurements led us to conclude that this transition is accompanied by the transfer of Trp residue to a more nonpolar environment. To support this conclusion, MD simulation studies were carried out.

As discussed in the Materials and Methods, MD simulations of both SAT1 and SAT3 were performed in the aqueous solution and also in the presence of urea. MD simulations help to gain atomistic insights of macromolecular systems and their dynamic behavior in various conditions such as mutational effects, ligand interaction, and the effects caused by destabilizers.[32,33] In his review of MD simulations of macromolecules, Martin Karplus, Noble laureate and credited with the first study published on protein folding dynamics in 1977, states the importance of MD simulation methods applied in the study of protein folding/unfolding.[32] In his view, MD simulations have the ability to provide intricate details of “individual particle motions as a function of time”, which is a significant observation to study the folding/unfolding of proteins. By studying the trajectories of atomic motions, we can estimate the folding/unfolding pathways and observe the changes occurring in the native structures of the proteins. The structural changes occurring in response to certain events are observed by calculating various geometrical and stereochemical properties of the macromolecules. Such properties include root-mean-square deviation (RMSD) of the atomic positions of the Cα or backbone atoms,[34] radius of gyration (Rg) of the proteins, changes in the atomic position and their effect on residual fluctuations in terms of root-mean-square fluctuation (RMSF), changes in the number of hydrogen bonds during the MD, perturbations in the solvent-accessible surface area (SASA) of the proteins, etc.

Urea-Induced Conformational Changes in SAT1 and SAT3

To observe the effects of higher concentrations of urea (4.0, 6.0, and 8.0 M) on SAT1 and SAT3, we performed MD simulations of each system for 100 ns in aqueous (0 M urea) and urea solutions. Time evolution plots of RMSD of all the systems (Figure A) suggest significant changes in the structure in comparison to the native conformations. Figure A shows RMSD values of Cα atoms of SAT1 with the following observations. (i) SAT1 in water shows relatively higher fluctuations in the Cα positions with a high drift up to 15 ns and then achieving balance for the rest of the simulation time. (ii) SAT1 in 4.0 M urea also shows an initial jump of 0.15 nm and then shows a drift of 0.17 nm and then reaches convergence after 40 ns of simulation time. Similarly, (iii) SAT1 in 6.0 M urea also shows an initial peak in the RMSD plot up to 10 ns then reaches convergence. (iv) SAT1 in 8.0 M urea shows similar trends to those in 4.0 and 6.0 M urea. These observations show that urea has a perturbing effect on the conformation of the SAT1 protein. The structure is significantly distorted in the effect of higher concentrations of urea. However, SAT1 in 6.0 and 8.0 M urea shows slightly less perturbance in comparison to SAT1 in a 4.0 M urea solution.

Figure 5

(A) RMSDs of Cα atoms are shown as a function of time for SAT1 in water (black), 4.0 M urea (red), 6.0 M urea (green), and 8.0 M urea (blue) at 300 K. (B) RMSDs of Cα atoms are shown as a function of time for SAT3 in water (black), 4.0 M urea (red), 6.0 M urea (green), and 8.0 M urea (blue) at 300 K.

An RMSD plot (Figure B) of SAT3 in water and urea at 4.0 and 6.0 M shows similar destabilizing effects on the structure of SAT3 in response to higher concentrations of urea. It was observed that SAT3 in 4.0 M urea solutions shows a high drift of around 0.27 nm up to 50 ns and then reaches convergence trending up to 100 ns. However, SAT3 in 6.0 and 8.0 M urea sees an initial jump of 0.2 nm and then shows a relatively stable pattern of RMSD fluctuation for the rest of the simulation time. The comparison of RMSD plots and average values of RMSD in Table of both SAT1 and SAT3 in 4.0, 6.0, and 8.0 M urea suggests that SAT1 is relatively more stable than SAT3 in solvent conditions.

Table 2

Average Values of Rg, RMSD, and RMSF for 100 ns of MD Simulations of SAT1 and SAT3 in Aqueous, 4.0 M Urea, 6.0 M Urea, and 8.0 M Urea

				RMSF (nm)
protein	system	Rg (nm)	RMSD (nm)	chain A	chain B	chain C
SAT1	native	2.587	0.243	0.124	0.136	0.130
	4.0 M urea	2.609	0.182	0.102	0.109	0.107
	6.0 M urea	2.631	0.166	0.114	0.108	0.107
	8.0 M urea	2.595	0.174	0.098	0.108	0.103
SAT3	native	2.581	0.261	0.160	0.135	0.135
	4.0 M urea	2.593	0.254	0.123	0.130	0.114
	6.0 M urea	2.590	0.210	0.107	0.109	0.102
	8.0 M urea	2.599	0.272	0.107	0.116	0.149

The radius of gyration (Rg) reflects the changes in the compactness of proteins during folding/unfolding,[35] hence it can be helpful to identify the changes in protein conformation during MD simulations. We calculated the Rg values of both SAT1 and SAT3 in the presence of 4.0 M, 6.0 M, and 8.0 M urea as a function of time. The R plot of SAT1 (Figure S5A) in 4.0, 6.0, and 8.0 M urea shows significant changes in the overall compactness of the protein. SAT1 in water (native state) shows a negative drift in the Rg up to 10 ns, then it reaches the convergence and follows the stable conformation up to 100 ns. On the other hand, SAT1 in 4.0 M, 6.0 M, and 8.0 M urea shows the open conformation of the protein, suggesting possible perturbation of the 3D structure of the SAT1. SAT1 in 6.0 M urea shows a higher perturbance in the structure up to 30 ns, then following a stable conformation up to 100 ns. We observed that in comparison to 4.0 and 6.0 M urea, 8.0 M urea has a more stabilizing effect on the structure of the SAT1. The similar trend is observed for SAT3 (Figure S5B) in a 4.0 M urea solution with comparatively lesser deviance than protein in a 6.0 M urea solution. However, for SAT3, 8.0 M urea is causing higher perturbance in comparison to 4.0 and 6.0 M urea, respectively.

Root mean square fluctuation (RMSF) is a helpful parameter to measure the conformational changes at the amino acid level. RMSF plots of both SAT1 and SAT3 show significant changes in the conformation of both the proteins in response to 4.0, 6.0, and 8.0 M urea solutions. Figure A shows higher fluctuations in the N-terminal segments of SAT1 followed by a slight stable conformation up to the 200th residue. However, there is a visible fluctuation in the atomic positions of amino acids of SAT1 for all of the monomeric units from the 200th to 225th residue. Similar observations are made for the SAT3 (Figure B) in higher concentrations of urea (4.0, 6.0, and 8.0 M). Hydrophobic tryptophan residues Trp195 and Trp295 in SAT1 and SAT3, respectively, show higher fluctuations in the RMSF plot (encircled region in Figure A and B), suggesting a relocation of Trp residues toward a more hydrophobic environment.

Figure 6

(A) RMSF of amino acid residues of SAT1 in water and different concentrations of urea. SAT1 in water (black), 4.0 M urea (red), 6.0 M urea (green), and 8.0 M urea (blue) at 300 K. Encircled region contains tryptophan 195 and 255 residues. (B) RMSF of amino acid residues of SAT3 in water and different concentrations of urea. SAT3 in water (black), 4.0 M urea (red), 6.0 M urea (green), and 8.0 M urea (blue) at 300 K. Encircled region contains tryptophan 195 and 255 residues.

Figures and 8 show the number of amino acid residues involved in the formation of secondary structure elements for SAT1 and SAT3 in aqueous and urea solutions. For both proteins, a secondary structure plot shows an increase in β sheets in 4.0, 6.0, and 8.0 M urea. Residues involved in β sheet formation are depicted with green color in Figure , where an increase in the number of residues involved in β sheet formation is observed.

Figure 7

Secondary structure plot of SAT1 in (A) aqueous solution, (B) 4.0 M urea, (C) 6.0 M urea, and (D) 8.0 M urea solutions.

Figure 8

Secondary structure plot of SAT3 in (A) aqueous, (B) 4.0 M urea, (C) 6.0 M urea, and (D) 8.0 M urea solutions.

The superimposed structures of the native SAT1 and SAT3 with the structures extracted from the 100 ns MD trajectories of SAT1 and SAT3 in 4.0, 6.0, and 8.0 M solutions along with secondary structure graphs generated by the STRIDE server (Figure S6 and S7) suggest an increase in β sheets and a decrease in α helical structure and coil. We also calculated the total solvent accessible surface area for SAT1 (Figure S8A) and SAT3 (Figure S8B), which indicates a significant increase in the SASA in response to 4.0, 6.0, and 8.0 M solution, suggesting an opening of the conformation which occurs in response to the opening of the native conformation. Free energy of solvation (Figure S9A and B) complements the changes in SASA of the SAT1 protein. We observed similar changes in the SASA of SAT3 for 4.0, 6.0, and 8.0 M urea, which is backed by the changes in the free energy of solvation with a significant increase in the values. Changes in the intramolecular hydrogen bonds are also helpful in identifying the structural changes during MD simulations. Hydrogen bond plots (Figures S10 and S11) of both SAT1 and SAT3 for 4.0, 6.0, and 8.0 M urea suggest a decrease in the number of the hydrogen bonds during MD simulation times, indicating possible disruption of the native structure of the protein. The average values of Rg, RMSD, and RMSF for 100 ns MD simulations of SAT1 and SAT3 in the native and urea-induced states are given in Table .

Conclusions

SAT isoforms are of great biological importance for E. histolytica for its growth and survival. However, the structural dynamics of SATs are still not studied well. Here, we have studied the effect of urea on the secondary and tertiary structure of SAT1 and SAT3 to understand its mechanism of folding as well as secondary structural transitions using molecular dynamics. We have observed that the conformational change in SAT1 and SAT3 isoforms in the presence of urea follows a two-state mechanism as revealed by normalized transition curves of all three optical properties [θ]222, F345, and Δε287. Furthermore, this conformational change involves α-helix → β-structure transition as revealed by [θ]222 measurements. These in vitro observations were complemented by 100 ns MD simulations of both SAT1 and SAT3 in the absence and presence of urea. Thus, these proteins which undergo an α-helix → β-sheet transition could be used as models to study neurodegenerative disease pathogenesis. SAT3 is a biologically more important protein, and its function is not regulated through a feedback mechanism of cysteine biosynthesis. E. histolytica could not survive if the SAT3 gene is knockdown, while the organism could survive if SAT1 and SAT2 genes are knockdown. Interestingly, SAT3 is less stable then SAT1, although it plays critical role in the survival of E. histolytica. Our studies suggested that the structural stability of SAT3 (SAT3 is approximately 2 kcal/mol less stable than SAT1) is compromised by nature, while attaining biologically important functions. The organism has evolved the SAT3 isoform to escape the feedback inhibition by the final product and to keep producing cysteine, but in the process SAT3 compromised its structural stability.