The Swedish Research Council (VR) is greatly acknowledged for financial support of this work by a young investigator grant #621-2013-5138. The PDC Center for High Performance Computing at the KTH Royal Institute of Technology is acknowledged for providing computational support.

1. In Silico Computer Modeling
<ol>
	<li>Download a PDB structure of the protein of interest from the protein data bank (PDB). Prepare the structure by removing excess subunits and then adding missing hydrogen atoms and explicit solvent using the &#34;Cell Neutralization and pKa prediction&#34; experiment in YASARA16. For membrane bound triterpene cyclases, suitable dimensions of the water box are 91x67x77 &#197;. Adjust the protonation state of the catalytically active amino acids manually. 
	Note: If necessary, a substrate analog in the PDB file can be modified to a real substrate by using the &#34;Edit|Swap|Atom&#34; command.</li>
	<li>Minimize the structure by the &#34;Energy Minimization&#34; command using the AMBER force field suite17. Use the particle mesh Ewald approach (PME)18 to account for long-range electrostatic interactions. Set the cut-off for van der Waals interactions according to standard settings. 
	Note: The &#34;Energy Minimization&#34; command removes conformational stress by a short steepest descent minimization, then a simulated annealing (timestep 2 fsec, atom velocities scaled down by 0.9 every 10th step) is performed until convergence is reached (i.e., improvement of energy by less than 0.012 kcal/mol per atom during 200 steps).</li>
</ol>
2. Model of Active Site Solvation and Water Access
<ol>
	<li>After the simulated annealing, run a 20 nsec molecular dynamics (MD) trajectory and generate at least 10 snapshots by the &#34;File|Save As|Simulation Snapshot&#34; command followed by &#34;Simulation On&#34;. Run the simulations on a standard computer with periodic boundary conditions under the canonical ensemble. Maintain the temperature at 298 K using a Berendsen thermostat.</li>
	<li>Prepare the snapshots obtained from 2.1 by deleting all water molecules and eventual ligands/substrates using the &#34;Edit|Delete|Residue&#34; command. Superpose each snapshot individually on the original PDB-structure by the &#34;Superpose|Object&#34; command to ensure that all structures share the same spatial position. Save each snapshot prepared in this way as a new PDB-file (using the &#34;File|Save as|PDB&#34; command). 
	Note: For experienced modelers: Write a script to automate this process according to the template given in Supplementary Code File 1.</li>
	<li>Use the prepared snapshots (PDB) from 2.2 that have been aligned in 3D-space as inputs to the software CAVER 3.019. For the analysis of a limited number of snapshots, run CAVER on a standard computer. Use a probe radius of 0.7 &#197;19 to assure the detection of tunnels that are specific for the transport of water molecules.</li>
	<li>Visualize the tunnels generated by CAVER 3.0 in a molecular graphics software. For easy visualization of tunnels, use the macro shown in Supplementary Code File 2.</li>
</ol>
3. In Silico Enzyme Engineering to Modify Water Patterns and Water Dynamics
<ol>
	<li>Identify the highest ranked tunnels according to the header &#34;ID&#34; listed in the output file &#34;summary.txt&#34; generated from the CAVER 3.0 software.</li>
	<li>Identify amino acids lining the highest ranked tunnels (3.1) and that display a small side chain pointing towards the channel interior by visual inspection (using the model obtained from 2.4). Identify single amino acid substitutions that will block the tunnel by increased steric hindrance using the &#34;Swap Residue&#34; command. Verify that the tunnel is obstructed by visual inspection, and then by repeating steps 1.2-3.1 of the protocol.</li>
</ol>
4. Expression of Mutated Genes
<ol>
	<li>Introduce mutations in the wild-type gene by Mutagenesis PCR using non-overlapping primers20.</li>
	<li>Add plasmid DNA containing the gene of interest to tubes with competent cells of a suitable cloning strain and incubate on ice for 30 min. Perform a heat shock transformation for 45 sec in a water bath with the temperature set to 42 &#176;C. Add 500 &#181;l of a fresh rich medium (2YT, 16 g/L tryptone, 10 g/L yeast extract, 5 g/L NaCl) and incubate at 37 &#176;C, 200 rpm for 45 min. Make a selection by plating the transformed cells on agar plates supplemented with appropriate antibiotics.</li>
	<li>After verifying the DNA sequence of the mutated gene, transform the plasmid DNA into an expression strain using heat shock as described above. For the expression of membrane bound triterpene cyclases, use BL21(DE3).</li>
	<li>Start a 5 ml O/N culture at 37 &#176;C of the expression strain in 2YT-media supplemented with the appropriate antibiotics.</li>
	<li>Inoculate a 2 L baffled shake flask, containing 300 ml 2YT-media supplemented with appropriate antibiotics, to a final OD of 0.05-0.09 (measured at 600 nm). After induction at an OD of 0.6-0.8 and protein expression, freeze the cell pellet and store at -80 &#176;C.
	<ol>
		<li>For membrane-bound triterpene cyclases in a pD861 vector, induce with 0.5 to 2 mM rhamnose and 4 hr of expression at 37 &#176;C to achieve high yields of protein21.</li>
	</ol>
	</li>
</ol>
5. Membrane Extraction Using a Normal Centrifuge and Protein Purification
<ol>
	<li>Prepare the following buffers: Resuspension buffer (100 mM potassium phosphate, pH 7.5 supplemented with Protease Inhibitor Tablets), Detergent buffer (1% (v/v) Triton X-100, 50 mM potassium phosphate, pH 7.5) and Reaction buffer (0.2% Triton X-100, 50 mM potassium phosphate, pH 7.5).
	<ol>
		<li>For squalene-hopene cyclases or other membrane-bound enzymes with a lower pH optimum, replace the potassium phosphate in the resuspension buffer with citrate. For such enzymes use the following buffers: Detergent buffer (1% Triton X-100, 60 mM citrate, pH 6.0) and Reaction buffer (0.2% Triton-X 100, 60 mM citrate, pH 6.0). 
		Note: If a His-tag is used for purification, supplement the resuspension buffer with 20 mM imidazole (final concentration).</li>
	</ol>
	</li>
	<li>Weigh a frozen cell pellet in a glass beaker. Resuspend the cells in resuspension buffer using a homogenizer to a final concentration of 0.3 g cells/ml. Lyse the resuspended cells by ultrasonication with an amplitude of 80% and a pulse of 1 sec on and 1 sec off. Use three repeating cycles of 50 sec each.</li>
	<li>Add detergent buffer to a final concentration of 0.2% (v/v) detergent. Equilibrate by end-over-end mixing at 4 &#176;C for 1 hr. Recover the membrane protein fraction by saving the supernatant after a single centrifugation step at 39,000 x g for 50 min at 4 &#176;C. 
	Note: If His-tag purification is used, add approximately 1.5 ml Ni-NTA agarose/g cells and incubate for 1 hr.</li>
	<li>Perform a first purification step by either ion exchange or His-tag purification21. Supplement the equilibration and elution buffers with 0.2% Triton X-100. Concentrate the purified protein five times using Centrifugal Filter Units with a cut-off of 10 kDa. 
	Note: The size of the Triton X-100 micelles results in a concentration of detergent to a final concentration of approximately 1%.</li>
	<li>Finalize purification by a gel filtration polishing step using the concentrated protein and a matrix compatible with a fractionation range for globular proteins of 2 &#215; 104-8 &#215; 106. Use the reaction buffer as running buffer. Measure the amount of purified protein by using the Bradford Ultra kit according to the manufacturer&#39;s protocol.</li>
</ol>
6. Kinetics
<ol>
	<li>Make a fresh 5 mM stock solution of substrate in reaction buffer. Obtain a homogenous emulsion by ultrasonication at 30% amplitude for 2-5 min.</li>
	<li>Equilibrate a thermomixer at the desired reaction temperature. Verify the temperature inside a glass vial containing water by an external thermometer.</li>
	<li>For single-substrate kinetics, dilute the emulsified substrate into at least five glass vials to the same final substrate concentration.
	<ol>
		<li>For one-pot relative kinetics with multiple substrates, prepare at least five identical glass vials by mixing all substrates in each vial. For hydrophobic triterpenes, use final substrate concentrations in the range of 10-200 &#181;M.</li>
	</ol>
	</li>
	<li>Preincubate the glass vials in the thermomixer for 10 min. Start the reaction by adding the enzyme. A total reaction volume of 1 ml is suitable. Use a shaking speed of at least 1,200 rpm.</li>
	<li>Stop the reactions at different time points by adding 500 &#181;l ethylacetate spiked with 100 &#181;M decanol as an integration standard.</li>
</ol>
7. Extraction and Thermodynamic Analysis
<ol>
	<li>Extract the reaction mixtures by vortexing and manually shaking the glass vials vigorously for 1 min. Centrifuge the glass vials at 9,600 x g in a table top centrifuge for 10 min at RT. Transfer the top layer (organic phase) to an empty tube. Add an additional 500 &#181;l of extraction solvent to the reaction tubes and repeat the extraction procedure.</li>
	<li>Dry the extracted reaction mixtures with Na2SO4. Vortex for 5 sec and let the tubes rest for 10 min. Perform a final centrifugation step at 9,600 x g for 1 min at RT using a table top centrifuge. Transfer the extracted samples to gas chromatography (GC) vials.</li>
	<li>Repeat the steps under 6.1-7.2 for each enzyme variant of interest, using at least four different initial concentration of the substrate ( for single substrate kinetics) and four different temperatures (for both single substrate and competitive kinetics).</li>
	<li>Perform GC-analysis as previously described3. Use a non-polar column for the analysis of hydrophobic pentacyclic products. Set the initial oven temperature to 120 &#176;C and use a temperature gradient of 5 &#176;C/min, depending on the products and the column.</li>
	<li>Integrate the peak areas of the product(s) and transform product peak areas into the corresponding product concentrations by utilizing the area of the integration standard (corresponding to 100 &#181;M). Plot the concentration of each product ([P]) versus the reaction time (t). Perform linear regression of the data points corresponding to less than 10% conversion. The initial reaction speed (V0) is given by the slope of the fitted line according to: 
	<img alt="Equation 1" src="/files/ftp_upload/53168/53168equation1.jpg" /> 
	Note: For one-pot relative kinetics with multiple substrates, V0 is the initial rate of a particular substrate under the influence of other substrates.</li>
	<li>For single substrate kinetics, plot V0/[E] against the corresponding substrate concentrations. The apparent kcat/KM value is given by the slope of the linear part of the graph according to: 
	<img alt="Equation 2" src="/files/ftp_upload/53168/53168equation2.jpg" /> 
	[S] is the concentration of substrate and [E] the concentration of enzyme used in the transformation. kcat/KM equates to the catalytic constant over the Michaelis constant. Repeat the analysis for each temperature and each enzyme variant.</li>
	<li>For single substrate kinetics, perform a thermodynamic analysis according to transition state theory22 
	<img alt="Equation 3" src="/files/ftp_upload/53168/53168equation3.jpg" /> 
	kb is the Boltzmann constant, h the Planck&#39;s constant, T the temperature in Kelvin, and R the gas constant. Perform a linear regression analysis of the plot of ln((kcat/KM)/((kb*T)/h))) versus 1/T. The activation enthalpy (&#916;H&#8225;) is given by (-slope)*R and the activation entropy (&#916;S&#8225;) is given by (intercept)*R. 
	Note: Likewise, an analysis under saturating substrate concentration can be performed by using kcat as rate constant in equation 3.</li>
	<li>For one-pot relative kinetics with multiple substrates, obtain the relative apparent 
	kcat/KM values from23: 
	(kcat/KM)A/(kcat/KM)B = V0,A/V0,B*[B]/[A] (4) 
	for which V0,A refers to the initial rate for substrate A in competition with substrate B.</li>
	<li>For one-pot relative kinetics with multiple substrates, obtain the relative thermodynamic parameters of activation for B, compared to A as reference, by nonlinear regression: 
	<img alt="Equation 5" src="/files/ftp_upload/53168/53168equation5.jpg" /></li>
	<li>Calculate the absolute activation parameters for substrate B from the activation enthalpy and entropy of reference compound A: 
	<img alt="Equation 6" src="/files/ftp_upload/53168/53168equation6.jpg" /></li>
</ol>

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The authors have nothing to disclose.

The most critical steps in achieving high quality experimental thermodynamical data for wild-type membrane enzyme and tunnel variants are: 1) generation of the computer model; 2) homogeneously purified proteins; 3) emulsified substrate stocks; 4) control of temperature during kinetics; 5) extraction of reaction mixtures using internal standard.
The generation of the computer model is greatly facilitated by the use of a software with a user-friendly interface that supports a variety of platforms. Hence, this protocol is premised on the YASARA modeling suite16 to make our strategy accessible even to modeling non-experts. A computer model for water tunnel identification should ideally be based on a crystal structure of the enzyme of interest24. For this purpose, the wealth of crystal structures available in the protein data bank is very beneficial. In our experience, a key aspect in the successful preparation of templates for tunnel identification is to keep crystallographic waters. It is of equal importance to use a solvated enzyme box when performing molecular dynamics simulations, which can be run on a normal computer. The triterpene cyclase from Alicyclobacillus acidocaldarius is stable in water during MD simulations3. However, keeping crystallographic detergents and/or using cell membrane mimics would perhaps be required for potentially unstable enzymes to allow for extended MD simulations. It is envisioned that the minimized crystal structure of highly challenging targets could provide important mechanistic insight using the protocol, although this would not capture dynamical aspects of tunnel organization.
CAVER19 in the basic mode, with a single or a limited number of snapshots as input, can be used by non-experts on a standard laptop. Based on our experience3, tunnels with a bottleneck radius (i.e., the radius at the most narrow point) smaller than 1 &#197; can be highly relevant for water, especially if crystallographic water molecules reside within the predicted tunnel (Figure 1, middle left). On the other hand, a larger bottleneck radius could imply a tunnel for the transport of the substrate in and out of the active site10. The script in Supplementary Code File 2 can be used by non-experts for the visualization of predicted tunnels. Future experiments will reveal whether homology models will be of sufficiently high resolution to allow for the atomistic study of water networks and dynamics. Shedding light on how performing molecular dynamics simulations, with and without a ligand present in the active site, influences the process of tunnel identification would also be of importance.
Kinetics of membrane proteins can constitute a formidable challenge25. The protocol herein is based on a simple membrane extraction protocol to obtain the membrane enzyme without the use of expensive equipment, such as an ultracentrifuge. The use of gel filtration as a final polishing step removes potential residual membrane particles and allows for defining an appropriate detergent environment25.
A key aspect in achieving reproducible kinetic results from the protocol is to emulsify the substrate stock solution by ultrasonication. Simple vortexing of hydrophobic substrates diluted in the reaction buffer gives inhomogeneous substrate-detergent mixtures. The pipetting of non-emulsified substrate solutions leads to irreproducible concentrations (confirmed by quantitative GC), which prevents accurate determination of initial rates. In contrast, the pipetting of properly emulsified stock solutions should result in linear regression analysis of initial rates with R2 in the range of 0.98-0.99. Another important aspect of hydrophobic substrates is the apparent substrate solubility and availability in the substrate-detergent mixtures. In fact, it was not possible to saturate the triterpene cyclase with the reference substrate squalene. For this reason apparent kcat/KM values are presented herein which could contain contributions from both binding and chemistry. However, it has been shown that chemistry is rate limiting for kcat/KM for the polycyclization cascade conducted by triterpene cyclases3.
It is of high importance to verify the actual temperature inside a reaction glass vial with an external thermometer. Still, linear fits can be poorer for variants with essential constant apparent kcat/KM values at different temperatures. This is stressed for the S168F variant herein (Figure 2A) with an activation enthalpy close to zero (Figure 2B and 2C). Very small temperature-dependent changes in apparent kcat/KM, could induce uncertainty in the observed activation entropy &#916;S&#8225; (i.e., the intercept in the linear plots in Figure 2B). In principle, the observed activation entropy could also be affected by a different abundance of active enzymes for different variants, which would not be detected by measuring the protein concentration. It is expected that experimental errors are reduced when mixing several substrates in one pot. This is because all the different substrates interact with the same amount of enzyme under these circumstances (equation 4). The use of an extraction solvent spiked with an internal standard is important to account for differences during extraction and/or GC-injection.
Transition state theory has been successfully used in enzymology26. This important theoretical framework was originally developed for unimolecular reactions in the gas phase. However, it has been shown that enzymes mainly work by lowering the classical activation energy barrier26. The transmission coefficient assumed to be one herein could affect the measured activation enthalpy and/or entropy. The contribution of tunneling, and other non-classical effects such as recrossing of the transition state, can roughly contribute 1,000-fold to catalysis26 corresponding to about 4 kcal/mol in energy. It can be seen that the activation entropy displayed by the wild-type enzyme (16 kcal/mol at 328 K, Figure 2C) is much larger than such non-classical effects caused by a non-uniform transmission coefficient. The impact of the transmission coefficient should decrease when comparing thermodynamic parameters of activation for wild type and tunnel variants using the protocol.
The Gibbs free energy of activation (&#916;G&#8225;) is composed of both an enthalpic (&#916;H&#8225;) and an entropic (-T*&#916;S&#8225;) term. Solvent reorganization in enzymes during catalysis can influence both parameters. The present protocol is expected to facilitate the study of these phenomena by assembling a toolbox of relevant and user friendly in silico computational tools with the necessary biophysical experimental framework. The method is envisaged to be useful for studying a plethora of enzymatic processes, including catalysis by membrane-bound enzymes.

Water constitutes a cornerstone for the chemistry of life1. Water patterns and solvation of enzyme active sites affect both the enthalpy and entropy of ligand binding1,2 and catalysis3 in a highly complex fashion extending beyond the hydrophobic effect2,3. NMR4, calorimetry2 and molecular modeling of solvated proteins have been used to shed light on the role of explicit water molecules in providing a driving force for ligand association5-8, specificity and activity2,9,10. Herein we present a unique methodology for the experimental assessment of the thermodynamical impact of solvent displacement on enzyme catalysis (Figure 1). Our combined strategy is based on using computer simulations in concert with enzyme engineering and thermodynamical analysis (Figure 1). This allows for shedding additional light on the impact of solvent dynamics on catalysis, which is currently poorly understood.
Stabilizing enthalpic interactions, provided by water-mediated hydrogen bonds in solvated enzyme active sites, can be offset by entropic penalties1. These entropic costs are associated with the decrease in the degrees of freedom displayed by water molecules confined within protein cavities, as compared to water in bulk5. The release of ordered water molecules can thus provide an entropic driving force for ligand association1 and catalysis3. A key aspect of solvent dynamics is the displacement of water molecules between the interior of proteins and the exterior solvent4. The accompanying changes in activation energy, enthalpy, and entropy11 are not completely understood on the molecular level. By obstructing individual tunnels responsible for the transport of water in enzymes, the importance of solvent dynamics and its contribution to the activation energy can be evaluated (Figure 1). Moreover, by performing one-pot kinetic experiments at different temperatures, the relative thermodynamical activation parameters of several substrates can be extracted from a reduced number of experiments (Figure 1, right). Our interdisciplinary method is validated for complex membrane-bound triterpene cyclase enzymes that generate polycyclic terpenes of high importance for life12. The protocol allows for the recovery of high amounts of membrane protein (10-20 mg/L) using a standard centrifuge.
Although enzymes evolved in water, the role of the solvent in promoting catalysis is usually neglected. In addition to protein dynamics13,14 that shape pre-organized active sites with electrostatic complementarity to the transition state15, water dynamics could be of high importance for efficient enzyme catalysis. By forging several interdisciplinary techniques, our aim is to facilitate the highly complex study of water dynamics and thermodynamics. Making these tools more accessible to the scientific community will lead to the development of new strategies in enzyme engineering and protein design for altered activities and specificities.

<table><tbody><tr><td>YASARA</td><td>YASARA Biosciences</td><td>http://www.yasara.org/</td><td>Molecular modeling and simulation program&amp;nbsp;</td></tr><tr><td>CAVER&amp;nbsp;</td><td>CaverSoft</td><td>http://caver.cz/</td><td>Tool for analysis of tunnels in proteins, free license for academic use</td></tr><tr><td>Bradford Ultra</td><td>Expedeon</td><td>BFU1L, BFU05L</td><td>Protein quantitation in solutions containing up to 1% detergent</td></tr><tr><td>Potter-Elvehjem homogenizer</td><td>VWR</td><td>432-0205, 432-0217</td><td>Homogenization of frozen cell pellet</td></tr><tr><td>Protease Inhibitor Cocktail Tablets</td><td>Roche</td><td>4693159001</td><td>Protease inhibitor</td></tr><tr><td>Centrifugal Filter Units</td><td>Millipore</td><td>UFC901008</td><td>Centrifugal filter units for the concentration of proteins, MWCO 10 kDa</td></tr><tr><td>Thermomixer&amp;nbsp;</td><td>Eppendorf</td><td>5382000015</td><td>Thermomixer for sample incubation</td></tr></tbody></table>

unraveling entropic rate acceleration induced by solvent dynamics in membrane enzymes

Enzyme catalysis evolved in an aqueous environment. The influence of solvent dynamics on catalysis is, however, currently poorly understood and usually neglected. The study of water dynamics in enzymes and the associated thermodynamical consequences is highly complex and has involved computer simulations, nuclear magnetic resonance (NMR) experiments, and calorimetry. Water tunnels that connect the active site with the surrounding solvent are key to solvent displacement and dynamics. The protocol herein allows for the engineering of these motifs for water transport, which affects specificity, activity and thermodynamics. By providing a biophysical framework founded on theory and experiments, the method presented herein can be used by researchers without previous expertise in computer modeling or biophysical chemistry. The method will advance our understanding of enzyme catalysis on the molecular level by measuring the enthalpic and entropic changes associated with catalysis by enzyme variants with obstructed water tunnels. The protocol can be used for the study of membrane-bound enzymes and other complex systems. This will enhance our understanding of the importance of solvent reorganization in catalysis as well as provide new catalytic strategies in protein design and engineering.

The importance of water dynamics in enzymatic polycyclization catalysis is implicated by in silico analysis and subsequent visualization of detected tunnels (using the script in Supplementary Code File 2). Following section 3 of the protocol, S168 is found to be a &#34;hot spot&#34; amino acid residue lining one of the tunnels in the triterpene cyclase enzyme from Alicyclobacillus acidocaldarius (Figure 1, middle left). By introducing the mutation S168F in silico, the tunnel is obstructed as seen by visual inspection (Figure 1, bottom left).
The reaction rate for the formation of polycyclic products displayed by the triterpene cyclase enzyme is highly sensitive to temperature (Figure 2A). Following the protocol (section 4-7), it is found that the apparent kcat/KM displayed by wild-type enzyme for the formation of pentacyclic products increases fiftyfold when the temperature is raised by 25 &#176;C (Figure 2A). Blocking individual tunnels by introducing a single point mutation has a significant effect on the experimentally determined absolute apparent kcat/KM values, as well as on their temperature dependence (Figure 2A).
From the experimentally determined kinetic parameters, linear thermodynamical plots are generated by equation 3 and by following section 6-7 of the protocol for single substrate kinetics (Figure 2B). The very large changes in activation entropy and enthalpy observed for variants harboring blocked tunnels implies a key role for water dynamics in promoting the polycyclization cascade (Figure 2C, calculations based on Figure 2B). Moreover, variants with altered water tunnels display an altered Gibbs free energy of activation (&#916;G&#8225;=&#916;H&#8225;-T*&#916;S&#8225;, Figure 2B-C). For instance, at 303 K the S168F tunnel variant displays a Gibbs free energy of activation of 14 kcal/mol compared to 16 kcal/mol for the wild-type enzyme.
Following the protocol in section 7, equation 4 allows for the calculation of apparent kcat/KM values for additional substrates from one-pot kinetic experiments (Figure 3A). Moreover, a linear thermodynamic plot (Figure 3B) can be constructed by running the competition essay at different temperatures (equation 5). In analogy to single substrate kinetics, the relative activation enthalpy and entropy for additional substrates compared to a reference compound are readily accessible from the intercept and slope of the linear fits to the experimental data. Absolute values of thermodynamic parameters of activation can be assessed from arithmetic addition by using thermodynamic data associated with the reference compound (equation 6). Using the protocol herein, generated results clearly show that membrane enzyme variants with blocked water tunnels display fundamentally different thermodynamic parameters of activation for substrates of different sizes (Figure 3C).
<img alt="Figure 1" src="/files/ftp_upload/53168/53168fig1.jpg" /> 
Figure 1. Protocol summary: in silico computer modeling in concert with experimental protein design and thermodynamic analysis allows for an enhanced understanding of the influence of solvent dynamics on enzyme catalysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53168/53168fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53168/53168fig2.jpg" /> 
Figure 2. Thermodynamic parameters of activation of wild-type and tunnel variants from single substrate kinetics. (A) Apparent kcat/KM values obtained from experimental data and by using equations 1 and 2. (B) Thermodynamical analysis using transition state theory and linear fit of the experimental data to equation 3. (C) Thermodynamic parameters of activation calculated from the plots in (B). The prefolded squalene substrate is shown with bonds formed/broken as dotted lines. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53168/53168fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53168/53168fig3.jpg" /> 
Figure 3. Thermodynamic parameters of activation of wild-type and tunnel variants for different substrates. (A) Relative kcat/KM values obtained from one-pot kinetics by using equation 4. (B) Thermodynamical plots of the kinetic data at different temperatures using equation 5. (C) Absolute thermodynamic parameters of activation calculated by equation 6 and by using the substrate squalene in Figure 2 as reference. Bonds formed/broken in the substrates are shown as dotted lines. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53168/53168fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Unraveling Entropic Rate Acceleration Induced by Solvent Dynamics in Membrane Enzymes at JoVE.com

Unraveling Entropic Rate Acceleration Induced by Solvent Dynamics in Membrane Enzymes

Channels for the transportation of water molecules in enzymes influence active site solvation and catalysis. Herein we present a protocol for the engineering of these additional catalytic motifs based on in silico computer modeling and experiments. This will enhance our understanding of the influence of solvent dynamics on enzyme catalysis.

Enzyme catalysis evolved in an aqueous environment. The influence of solvent dynamics on catalysis is, however, currently poorly understood and usually ...

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8.9K Views.  KTH Royal Institute of Technology. Channels for the transportation of water molecules in enzymes influence active site solvation and catalysis. Herein we present a protocol for the engineering of these additional catalytic motifs based on in silico computer modeling and experiments. This will enhance our understanding of the influence of solvent dynamics on enzyme catalysis.

Video: Unraveling Entropic Rate Acceleration Induced by Solvent Dynamics in Membrane Enzymes

Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

Membrane protein folding is an emerging topic with both fundamental and health-related significance. The abundance of membrane proteins in cells underlies the need for comprehensive study of the folding of this ubiquitous family of proteins. Additionally, advances in our ability to characterize diseases associated with misfolded proteins have motivated significant experimental and theoretical efforts in the field of protein folding. Rapid progress in this important field is unfortunately hindered by the inherent challenges associated with membrane proteins and the complexity of the folding mechanism. Here, we outline an experimental procedure for measuring the thermodynamic property of the Gibbs free energy of unfolding in the absence of denaturant, &Delta;G&deg;H2O, for a representative integral membrane protein from E. coli. This protocol focuses on the application of fluorescence spectroscopy to determine equilibrium populations of folded and unfolded states as a function of denaturant concentration. Experimental considerations for the preparation of synthetic lipid vesicles as well as key steps in the data analysis procedure are highlighted. This technique is versatile and may be pursued with different types of denaturant, including temperature and pH, as well as in various folding environments of lipids and micelles. The current protocol is one that can be generalized to any membrane or soluble protein that meets the set of criteria discussed below.

This video article details the experimental procedure for obtaining the Gibbs free energy of membrane protein folding by tryptophan fluorescence.

Membrane protein folding is an emerging topic with both fundamental and health-related significance.  The abundance of membrane proteins in cells ...

Bioengineering

Sedimentation Equilibrium of a Small Oligomer-forming Membrane Protein: Effect of Histidine Protonation on Pentameric Stability

Analytical ultracentrifugation (AUC) can be used to study reversible interactions between macromolecules over a wide range of interaction strengths and under physiological conditions. This makes AUC a method of choice to quantitatively assess stoichiometry and thermodynamics of homo- and hetero-association that are transient and reversible in biochemical processes. In the modality of sedimentation equilibrium (SE), a balance between diffusion and sedimentation provides a profile as a function of radial distance that depends on a specific association model. Herein, a detailed SE protocol is described to determine the size and monomer-monomer association energy of a small membrane protein oligomer using an analytical ultracentrifuge. AUC-ES is label-free, only based on physical principles, and can be used on both water soluble and membrane proteins. An example is shown of the latter, the small hydrophobic (SH) protein in the human respiratory syncytial virus (hRSV), a 65-amino acid polypeptide with a single &#945;-helical transmembrane (TM) domain that forms pentameric ion channels. NMR-based structural data shows that SH protein has two protonatable His residues in its transmembrane domain that are oriented facing the lumen of the channel. SE experiments have been designed to determine how pH affects association constant and the oligomeric size of SH protein. While the pentameric form was preserved in all cases, its association constant was reduced at low pH. These data are in agreement with a similar pH dependency observed for SH channel activity, consistent with a lumenal orientation of the two His residues in SH protein. The latter may experience electrostatic repulsion and reduced oligomer stability at low pH. In summary, this method is applicable whenever quantitative information on subtle protein-protein association changes in physiological conditions have to be measured.&#160;&#160;

Sedimentation equilibrium (SE) can be used to study protein-protein interactions in a physiological environment. This manuscript describes the use of this technique to determine the effect of pH on the stability of a homo-pentamer formed by the small hydrophobic (SH) protein encoded by the human syncytial respiratory virus (hRSV).

Analytical ultracentrifugation (AUC) can be used to study reversible interactions between macromolecules over a wide range of interaction strengths and ...

Detergent-free Ultrafast Reconstitution of Membrane Proteins into Lipid Bilayers Using Fusogenic Complementary-charged Proteoliposomes.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers are less compatible with detergents.
Here we describe a strategy for bypassing this fundamental limitation using fusogenic oppositely charged liposomes bearing a membrane protein of interest. Fusion between such vesicles occurs within 5 min in a low ionic strength buffer. Positively charged fusogenic liposomes can be used as simple shuttle vectors for detergent-free delivery of membrane proteins into biomimetic target lipid bilayers, which are negatively charged. We also show how to reconstitute membrane proteins into fusogenic proteoliposomes with a fast 30-min protocol.
Combining these two approaches, we demonstrate a fast assembly of an electron transport chain consisting of two membrane proteins from E. coli, a primary proton pump bo3-oxidase and F1Fo ATP synthase, in membranes of vesicles of various sizes, ranging from 0.1 to &#62;10 microns, as well as ATP production by this chain.

Here we present two ultrafast protocols for reconstitution of membrane proteins into fusogenic proteoliposomes, and fusion of such proteoliposomes with target lipid bilayers for detergent-free delivery of these membrane proteins into the postfusion bilayer. The combination of these approaches enables fast and easily controlled assembly of complex multi-component membrane systems.

Detergents are indispensable for delivery of membrane proteins into 30-100 nm small unilamellar vesicles, while more complex, larger model lipid bilayers ...

Biochemistry

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used for ATP production. The amphiphilic nature of membrane proteins requires particular attention to their handling, and reconstitution into the natural lipid environment is indispensable when studying membrane transport processes like proton translocation. Here, we detail a method that has been used for the investigation of the proton-pumping mechanism of membrane redox enzymes, taking cytochrome bo3 from Escherichia coli as an example. A combination of electrochemistry and fluorescence microscopy is used to control the redox state of the quinone pool and monitor pH changes in the lumen. Due to the spatial resolution of fluorescent microscopy, hundreds of liposomes can be measured simultaneously while the enzyme content can be scaled down to a single enzyme or transporter per liposome. The respective single enzyme analysis can reveal patterns in the enzyme functional dynamics that might be otherwise hidden by the behavior of the whole population. We include a description of a script for automated image analysis.

Here, we present a protocol to study the molecular mechanism of proton translocation across lipid membranes of single liposomes, using cytochrome bo3 as an example. Combining electrochemistry and fluorescence microscopy, pH changes in the lumen of single vesicles, containing single or multiple enzyme, can be detected and analyzed individually.

Proton-pumping enzymes of electron transfer chains couple redox reactions to proton translocation across the membrane, creating a proton-motive force used ...

A Model Membrane Platform for Reconstituting Mitochondrial Membrane Dynamics

Mitochondrial dynamics is essential for the organelle&#8217;s diverse functions and cellular responses. The crowded, spatially complex, mitochondrial membrane is a challenging environment to distinguish regulatory factors. Experimental control of protein and lipid components can help answer specific questions of regulation. Yet, quantitative manipulation of these factors is challenging in cellular assays. To investigate the molecular mechanism of mitochondria inner-membrane fusion, we introduced an in vitro reconstitution platform that mimics the lipid environment of the mitochondrial inner-membrane. Here we describe detailed steps for preparing lipid bilayers and reconstituting mitochondrial membrane proteins. The platform allowed analysis of intermediates in mitochondrial inner-membrane fusion, and the kinetics for individual transitions, in a quantitative manner. This protocol describes the fabrication of bilayers with asymmetric lipid composition and describes general considerations for reconstituting transmembrane proteins into a cushioned bilayer. The method may be applied to study other membrane systems.

Mitochondrial fusion is an important homeostatic reaction underlying mitochondrial dynamics. Described here is an in vitro reconstitution system to study mitochondrial inner-membrane fusion that can resolve membrane tethering, docking, hemifusion, and pore opening. The versatility of this approach in exploring cell membrane systems is discussed.

Mitochondrial dynamics is essential for the organelle&#8217;s diverse functions and cellular responses. The crowded, spatially complex, mitochondrial ...

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

Most human body proteins' activity and functionality are related to configurational changes of entire subdomains within the protein crystal structure. The crystal structures build the basis for any calculation that describes the structure or dynamics of a protein, most of the time with strong geometrical restrictions. However, these restrictions from the crystal structure are not present in the solution. The structure of the proteins in the solution may differ from the crystal due to rearrangements of loops or subdomains on the pico to nanosecond time scale (i.e., the internal protein dynamics time regime). The present work describes how slow motions on timescales of several tens of nanoseconds can be accessed using neutron scattering. In particular, the dynamical characterization of two major human proteins, an intrinsically disordered protein that lacks a well-defined secondary structure and a classical antibody protein, is addressed by neutron spin echo spectroscopy (NSE) combined with a wide range of laboratory characterization methods. Further insights into protein domain dynamics were achieved using mathematical modeling to describe the experimental neutron data and determine the crossover between combined diffusive and internal protein motions. The extraction of the internal dynamic contribution to the intermediate scattering function obtained from NSE, including the timescale of the various movements, allows further vision into the mechanical properties of single proteins and the softness of proteins in their nearly natural environment in the crowded protein solution.

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

The present protocol describes methods for investigating the structure and dynamics of two model proteins that have an important role in human health. The technique combines bench-top biophysical characterization with neutron spin echo spectroscopy to access the dynamics at time and length scales relevant for protein interdomain motions.

Most human body proteins' activity and functionality are related to configurational changes of entire subdomains within the protein crystal structure. The ...

Engineering

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without labeling other than deuterium. In particular, neutron backscattering spectroscopy records the scattering signals at multiple scattering angles simultaneously and is well suited to study the dynamics of biological systems on the ps-ns timescale. By employing D2O-and possibly deuterated buffer components-the method allows monitoring of both center-of-mass diffusion and backbone and side-chain motions (internal dynamics) of proteins in liquid state.
Additionally, hydration water dynamics can be studied by employing powders of perdeuterated proteins hydrated with H2O. This paper presents the workflow employed on the instrument IN16B at the Institut Laue-Langevin (ILL) to investigate protein and hydration water dynamics. The preparation of solution samples and hydrated protein powder samples using vapor exchange is explained. The data analysis procedure for both protein and hydration water dynamics is described for different types of datasets (quasielastic spectra or fixed-window scans) that can be obtained on a neutron backscattering spectrometer.
The method is illustrated with two studies involving amyloid proteins. The aggregation of lysozyme into &#181;m sized spherical aggregates-denoted particulates-is shown to occur in a one-step process on the space and time range probed on IN16B, while the internal dynamics remains unchanged. Further, the dynamics of hydration water of tau was studied on hydrated powders of perdeuterated protein. It is shown that translational motions of water are activated upon the formation of amyloid fibers. Finally, critical steps in the protocol are discussed as to how neutron scattering is positioned regarding the study of dynamics with respect to other experimental biophysical methods.

Neutron backscattering spectroscopy offers a nondestructive and label-free access to the ps-ns dynamics of proteins and their hydration water. The workflow is presented with two studies on amyloid proteins: on the time-resolved dynamics of lysozyme during aggregation and on the hydration water dynamics of tau upon fiber formation.

Neutron scattering offers the possibility to probe the dynamics within samples for a wide range of energies in a nondestructive manner and without ...

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient association of pore-forming and auxiliary subunits, lateral diffusion, and clustering with other proteins. However, the relationship between lateral diffusion and function is poorly understood. To approach this problem, we describe how lateral mobility and activity of individual channels in supported lipid membranes can be monitored and correlated using total internal reflection fluorescence (TIRF) microscopy. Membranes are fabricated on an ultrathin hydrogel substrate using the droplet interface bilayer (DIB) technique. Compared to other types of model membranes, these membranes have the advantage of being mechanically robust and suitable for highly sensitive analytical techniques. This protocol measures Ca2+ ion flux through single channels by observing the fluorescence emission of a Ca2+-sensitive dye in close proximity to the membrane. In contrast to classical single-molecule tracking approaches, no fluorescent fusion proteins or labels, which can interfere with lateral movement and function in the membrane, are required. Possible changes in ion flux associated with conformational changes of the protein are only due to protein lateral motion in the membrane. Representative results are shown using the mitochondrial protein translocation channel TOM-CC and the bacterial channel OmpF. In contrast to OmpF, the gating of TOM-CC is very sensitive to molecular confinement and the nature of lateral diffusion. Hence, supported droplet-interface bilayers are a powerful tool to characterize the link between lateral diffusion and the function of ion channels.

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence TIRF Microscopy

This protocol describes how to use TIRF microscopy to track individual ion channels and determine their activity in supported lipid membranes, thereby defining the interplay between lateral membrane movement and channel function. It describes how to prepare the membranes, record the data, and analyze the results.

High-resolution imaging techniques have shown that many ion channels are not static, but subject to highly dynamic processes, including the transient ...

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This feature is attributed to the mechanical and dynamic properties of the cell cytoskeleton, notably, the actomyosin cytoskeleton, which is an active gel of polar actin filaments, myosin motors, and accessory proteins that exhibit intrinsic contraction properties. The usually accepted view is that the cytoskeleton behaves as a viscoelastic material. However, this model cannot always explain the experimental results, which are more consistent with a picture describing the cytoskeleton as a poroelastic active material-an elastic network embedded with cytosol. Contractility gradients generated by the myosin motors drive the flow of the cytosol across the gel pores, which infers that the mechanics of the cytoskeleton and the cytosol are tightly coupled. One main feature of poroelasticity is the diffusive relaxation of stresses in the network, characterized by an effective diffusion constant that depends on the gel elastic modulus, porosity, and cytosol (solvent) viscosity. As cells have many ways to regulate their structure and material properties, our current understanding of how cytoskeleton mechanics and cytosol flow dynamics are coupled remains poorly understood. Here, an in vitro reconstitution approach is employed to characterize the material properties of poroelastic actomyosin gels as a model system for the cell cytoskeleton. Gel contraction is driven by myosin motor contractility, which leads to the emergence of a flow of the penetrating solvent. The paper describes how to prepare these gels and run experiments. We also discuss how to measure and analyze the solvent flow and gel contraction both at the local and global scales. The various scaling relations used for data quantification are given. Finally, the experimental challenges and common pitfalls are discussed, including their relevance to cell cytoskeleton mechanics.

The Mechanics of Poro-Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

In this work, an in vitro reconstitution approach is employed to study the poroelasticity of actomyosin gels under controlled conditions. The dynamics of the actomyosin gel and the embedded solvent are quantified, through which the network poroelasticity is demonstrated. We also discuss the experimental challenges, common pitfalls, and relevance to cell cytoskeleton mechanics.

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This ...

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral budding, among other vital cellular processes. For efficient biological activity, cell membranes should be rigid enough to maintain the integrity of the cell and its compartments yet fluid enough to allow membrane components, such as proteins and functional domains, to diffuse and interact. This delicate balance of elastic and fluid membrane properties, and their impact on biological function, necessitate a better understanding of collective membrane dynamics over mesoscopic length and time scales of key biological processes, e.g., membrane deformations and protein binding events. Among the techniques that can effectively probe this dynamic range is neutron spin echo (NSE) spectroscopy. Combined with deuterium labeling, NSE can be used to directly access bending and thickness fluctuations as well as mesoscopic dynamics of select membrane features. This paper provides a brief description of the NSE technique and outlines the procedures for performing NSE experiments on liposomal membranes, including details of sample preparation and deuteration schemes, along with instructions for data collection and reduction. The paper also introduces data analysis methods used to extract key membrane parameters, such as the bending rigidity modulus, area compressibility modulus, and in-plane viscosity. To illustrate the biological importance of NSE studies, select examples of membrane phenomena probed by NSE are discussed, namely, the effect of additives on membrane bending rigidity, the impact of domain formation on membrane fluctuations, and the dynamic signature of membrane-protein interactions.

This paper describes the protocols for sample preparation, data reduction, and data analysis in neutron spin echo (NSE) studies of lipid membranes. Judicious deuterium labeling of lipids enables access to different membrane dynamics on mesoscopic length and time scales, over which vital biological processes occur.

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral ...

Unraveling Entropic Rate Acceleration Induced by Solvent Dynamics in Membrane Enzymes

Summary

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Chapters in this video

Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

Sedimentation Equilibrium of a Small Oligomer-forming Membrane Protein: Effect of Histidine Protonation on Pentameric Stability

Detergent-free Ultrafast Reconstitution of Membrane Proteins into Lipid Bilayers Using Fusogenic Complementary-charged Proteoliposomes.

Single Liposome Measurements for the Study of Proton-Pumping Membrane Enzymes Using Electrochemistry and Fluorescent Microscopy

A Model Membrane Platform for Reconstituting Mitochondrial Membrane Dynamics

Study of Protein Dynamics via Neutron Spin Echo Spectroscopy

High-Resolution Neutron Spectroscopy to Study Picosecond-Nanosecond Dynamics of Proteins and Hydration Water

Single-Molecule Imaging of Lateral Mobility and Ion Channel Activity in Lipid Bilayers using Total Internal Reflection Fluorescence (TIRF) Microscopy

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions