Using NMR Titration Experiments to study E. coli FAS-II and AcpP Mediated Protein-Protein Interactions

Abstract

Acyl carrier proteins (ACPs) are central to many primary and secondary metabolic pathways. In E. coli fatty acid biosynthesis (FAB), the central ACP, AcpP, transports intermediates to a suite of partner proteins (PP) for iterative modification and elongation. The regulatory protein-protein interactions that occur between AcpP and the PP in FAB is poorly understood due to the dynamic and transient nature of these interactions. Solution state NMR spectroscopy can reveal information at the atomic level through experiments such as the 2D heteronuclear single quantum coherence (HSQC). The following protocol describes NMR HSQC titration experiments that can elucidate biomolecular recognition events.

1. Introduction

Protein-protein interactions (PPIs) regulate essential biosynthetic pathways such as fatty acid biosynthesis, polyketide synthesis, and non-ribosomal peptide synthesis. [1,2] These pathways are carrier protein dependent, as they rely on a central carrier protein that shuttles a covalently bound substrate to various partner proteins (PPs) for modification and elongation. The PPIs between carrier protein’s and the PPs in their respective pathways are crucial for proper function and product formation, however the transient and dynamic nature of these interactions has left the molecular basis of these recognition events poorly understood. Here we will focus on the application of solution-phase protein NMR titration as a tool to provide restraints for in silico docking analysis, with a demonstration of these techniques upon fatty acid biosynthesis (FAB) in Escherichia coli.

FAB in E. coli has been extensively studied and these studies have provided much of the foundational knowledge for bacterial FAB [3]. FAB occurs via two distinct organizations. Eukaryotes predominantly use the modular type I fatty acid synthase (FAS-I) which contains all proteins as domains on a large megasynthase. Typically, bacteria, such as E. coli, deploy the type II fatty acid synthase (FAS-II) which is comprised of discrete individual proteins that iteratively interact with a central carrier protein bearing a substrate or intermediate (Figure 1).

Figure 1:

Diagram of E. coli fatty acid biosynthesis (FAS-II) highlighting the varied PPI that are mediated by ACPs. In E.coi FAS-II, the ACP will iteratively interact with a suite of PP to produce the final product.

In E. coli FAS-II the central acyl carrier protein (ACP), AcpP is the ACP responsible for transporting the growing substrate to each of the PP in the pathway for iterative modification and elongation [4]. A key feature of AcpP is its ability to sequester the substrate into the hydrophobic pocket to prevent premature hydrolysis. When AcpP comes into the appropriate proximity of a PP, the substrate undergoes “chain-flipping” into the pocket of the PP which has been found to be more hydrophobic than the pocket of AcpP. After the substrate has been modified, it is resequestered back into the AcpP pocket [5–7]. In addition to “chain-flipping”, AcpP and the PP in FAS-II must orchestrate a series of complex events, and the full spectrum of the PPi in these events remains elusive. NMR Titration studies have provided valuable insight into the biomolecular recognition in E. coli PPi [8–13], but further studies are needed to fully understand the complete choreography.

NMR titration experiments have been validated for studying protein-small molecule and protein-ligand interactions. Additional methods can be used to study binding and kinetics, but NMR titration experiments elucidate PPI at the atomic level providing a more in-depth, comprehensive analysis.[14–15] In conventional NMR titration experiments, the concentration of the receptor protein is held constant, and the ligand or small molecule concentration is steadily altered. This gradient alteration reveals the dynamic changes that occur at the surface residues of the receptor protein that are induced by the ligand or small molecule. These changes are made apparent through the mapping of complexation induced changes in the chemical environment of the proteins, also known as chemical shift perturbations (CSPs). By mapping the CSP, the locations of binding sites, allosteric regulation, surface interactions, and residue specific information can be revealed [16]. In ACP mediated interactions, the ACP carrier shuttles the substrate that is covalently attached to a conserved serine residue to the various PP in the pathway. In these interactions, the PP functions as the receptor and the ACP functions as the ligand. The provided method is reversed from traditional methods and holds the AcpP concentration constant while varying the PP concentration. This change in convention allows a refined focus that elucidates AcpP residue specific information that has been previously inaccessible.

2. Materials and Methods for Proteins

2.1. M9 Minimal Media

Isotopically labeled proteins are grown in minimal media with ¹⁵N and/or ¹³C isotopes added to control the carbon and nitrogen sources. This protocol for M9 minimal media with ¹⁵N is as follows:

1. Weigh out the following reagents:

   1. 6g Na 6g Na₂HPO₄
   2. 3g KH₂PO₄
   3. 0.5g NaCl
   4. 0.25g MgSO₄*7H₂O
   5. 0.015g CaCl₂*2H₂O
   6. 0.1g thiamine
   7. 0.003g FeSO₄*7H₂O
   8. 8g D-glucose
   9. 1g ¹⁵N-NH₄Cl

2. Dissolve all reagents in 1L of DI water

3. Sterile filter the prepared media into a clean, dry, autoclaved reagent bottle with a lid.

2.2. Lysis Buffer

1. Weigh out the following reagents

   1. 6.057 g Tris
   2. 8.766 g NaCl
   3. 100mL glycerol

2. Add all reagents to 500mL of ultra-pure water in a 1L reagent bottle

3. Adjust the pH to 7.4

4. Fill to 1L with ultra-pure water

5. Final composition is 50mM Tris, 150mM NaCl, 10% glycerol, pH 7.4

2.3. Urea-PAGE (Polyacrylamide gel electrophoresis)

Urea-PAGE is a conformationally sensitive electrophoresis method that allows the resolution of proteins that exist in a multitude of states. This method is particularly useful for monitoring the substrate modification of carrier proteins.

1. Set up a gel casting stand such as the Mini-PROTEAN Tetra Cell Casting Stand

2. Prepare 20% Urea Gels

   1. Prepare Resolving layer

      1. In a 15mL falcon tube, combine the following:

         1. 3.1 mL 10 M Urea
         2. 6.1 mL 40% acrylamide
         3. 3.2 mL 1.5M Tris, pH 8.8
         4. 0.125 mL ice cold 10% ammonium persulfate (APS) w/v
         5. 0.005 mL N, N, N, N’-Tetramethyl-ethylenediamine (TEMED)
      2. After all reagents are combined, invert the falcon tube 2–3 times to thoroughly mix. Quickly pour into the casting gel plates up to approximately 1.5 cm from the top.
      3. Fill the remaining 1.5 cm with isopropanol (IPA)
      4. Allow to polymerize for 45 minutes or until set. **There will be a small remainder in the falcon tube. When the remainder has polymerized, the gels will be set**
      5. After polymerization, decant off the IPA layer
   2. Prepare the Stacking layer

      1. In a 15mL falcon tube, combine the following:

         1. 2.9 mL ultra-pure water
         2. 0.5 mL 40% acrylamide
         3. 0.5 mL 1.0M Tris, pH 6.8
         4. 0.04 mL 10% APS
         5. 0.005 mL TEMED
      2. After all reagents are combined, invert the falcon tube 2–3 times to thoroughly mix. Quickly pour on top of the set resolving layer and fill to the top.
      3. Insert combs with desired number of lanes
      4. Allow to polymerize for 45 minutes or until set. **There will be a small remainder in the falcon tube. When the remainder has polymerized, the gels will be set**
   3. Gels can be stored in urea buffer, lying flat, at 4°C
3. Prepare 1L of 10X Urea Running Buffer

   1. Fill 1L bottle with 700mL of ultra-pure water
   2. Add 30.3g Tris
   3. Add 144.10g glycine. **add in batches while using a magnetic spin bar to allow faster solubilization**
   4. Fill to 1L with ultra-pure water
   5. Final composition 250mM Tris, 1.92M glycine, pH 8.3 **pH will naturally be between 8.3–8.9. Do not adjust pH**
4. Prepare Samples

   1. Prepare 5X Urea Loading Dye **Must be stored at −20°C due to DTT. If DTT is omitted, stock solution can be stored at room temperature and DTT added when preparing the individual samples**

      1. 250mM Tris pH 6.8
      2. 0.5% Bromophenol blue
      3. 500mM DTT
      4. 50% glycerol
   2. Prepare samples

      1. For each sample prepare a total volume of 15uL

         1. 3uL 5X Urea Loading dye
         2. 12uL of sample **For more pure protein samples, ~2ug of protein is ideal per well. For less pure samples ~20ug of protein is ideal. If the protein sample is purer, reduce volume of protein sample as necessary and fill to 12uL with lysis buffer**
5. Running the gel

   1. Set up a 1-D vertical gel electrophoresis system such as the Bio-Rad Mini-PROTEAN Tetra Cell
   2. Secure the 20% urea gel and plastic dummy plate in the electrode assembly plate.
   3. Fill the electrode assembly with 1X urea running buffer and remove the comb from the gel
   4. Load 8–15uL of sample into a well. Repeat for all samples.
   5. Fill the chamber with 1X urea buffer to the designated fill line and top off the chamber of the electrode assembly as needed if the volume was decreased during sample loading.
   6. Run at 200V for 120–150 minutes. **For urea-gels, the dye front will run completely off the gel. Do not stop to prevent the dye front from running off or there will not be sufficient resolution**
6. Visualizing the gel

   1. When the run is complete, carefully remove the gel from the gel cassette and transfer to a container. Rinse lightly with DI water
   2. “Fix” the gel by soaking in a solution of 10% glacial acetic acid, 50% methanol, and 40% water for 30 minutes to overnight.
   3. Decant off the “fix” solution and rinse lightly with DI water
   4. Stain 30 minutes to overnight in Coomassie Brilliant Blue
   5. Decant off Coomassie Brilliant Blue and rinse with DI water
   6. Soak in DI water with trace amounts of ethanol to remove background staining. If gel is over stained, repeat step b and stain again for a shorter duration

2.4. SDS-PAGE (Sodium Dodecyl Sulfate Polyacrylamide gel electrophoresis)

1. Set up a gel casting stand such as the Mini-PROTEAN Tetra Cell Casting Stand

2. Prepare 12% SDS Gels

   1. Prepare Resolving layer

      1. In a 15mL falcon tube, combine the following:

         1. 3.7 mL 40% acrylamide
         2. 3.1 mL 1.5M Tris, pH 8.8
         3. 5.4 mL ultra pure water
         4. 0.120 uL 10% SDS
         5. 0.120 mL ice cold 10% ammonium persulfate (APS) w/v
         6. 0.005 mL N, N, N, N’-Tetramethyl-ethylenediamine (TEMED)
      2. After all reagents are combined, invert the falcon tube 2–3 times to thoroughly mix. Quickly pour into the casting gel plates up to approximately 1.5 cm from the top.
      3. Fill the remaining 1.5 cm with isopropanol (IPA)
      4. Allow to polymerize for 45 minutes or until set. **There will be a small remainder in the falcon tube. When the remainder has polymerized, the gels will be set**
      5. After polymerization, decant off the IPA layer
   2. Prepare the Stacking layer

      1. In a 15mL falcon tube, combine the following:

         1. 2.9 mL ultra-pure water
         2. 0.5 mL 40% acrylamide
         3. 0.5 mL 1.0M Tris, pH 6.8
         4. 0.04 mL 10% SDS
         5. 0.04 mL 10% APS
         6. 0.005 mL TEMED
      2. After all reagents are combined, invert the falcon tube 2–3 times to thoroughly mix. Quickly pour on top of the set resolving layer and fill to the top.
      3. Insert combs with desired number of lanes
      4. Allow to polymerize for 45 minutes or until set. **There will be a small remainder in the falcon tube. When the remainder has polymerized, the gels will be set**
   3. Gels can be stored in SDS buffer, lying flat, at 4°C
3. Prepare 1L of 10X SDS Running Buffer

   1. Fill 1L bottle with 700mL of ultra-pure water
   2. Add 30.3g Tris
   3. Add 144.10g glycine. **add in batches while using a magnetic spin bar to allow faster solubilization**
   4. Add 10g SDS
   5. Fill to 1L with ultra-pure water
   6. Final composition 250mM Tris, 1.92M glycine, pH 8.3 **pH will naturally be between 8.3–8.9. Do not adjust pH**
4. Sample preparation

   1. Prepare 5X SDS Loading Dye **Must be stored at −20°C due to DTT. If DTT is omitted, stock solution can be stored at room temperature and DTT added when preparing the individual samples**

      1. 10% SDS
      2. 250mM Tris pH 6.8
      3. 0.5% Bromophenol blue
      4. 500mM DTT
      5. 50% glycerol
   2. Prepare samples

      1. For each sample prepare a total volume of 15uL

         1. 3uL 5X SDS Loading dye
         2. 12uL of sample **For more pure protein samples, ~2ug of protein is ideal per well. For less pure samples ~20ug of protein is ideal. If the protein sample is purer, reduce volume of protein sample as necessary and fill to 12uL with lysis buffer**
         3. Heat samples at 90–100°C for 5 minutes
         4. Remove from heat and proceed to “Running the gel”
5. Running the gel

   1. Set up a 1-D vertical gel electrophoresis system such as the Bio-Rad Mini-PROTEAN Tetra Cell
   2. Secure the 12% SDS gel and plastic dummy plate in the electrode assembly plate.
   3. Fill the electrode assembly with 1X SDS running buffer and remove the comb from the gel
   4. Load 8–15uL of appropriate protein ladder into the first lane
   5. Load 8–15uL of sample into a subsequent well. Repeat for all samples.
   6. Fill the chamber with 1X SDS buffer to the designated fill line and top off the chamber of the electrode assembly as needed if the volume was decreased during sample loading.
   7. Run at 200V for 30–45 minutes. **Do not allow the dye front to run off the gel. Watch carefully and stop the run when the dye front is 0.5–1 cm from the base of the gel**
6. Visualizing the gel

   1. When the run is complete, carefully remove the gel from the gel cassette and transfer to a container. Rinse lightly with DI water
   2. “Fix” the gel by soaking in a solution of 10% glacial acetic acid, 50% methanol, and 40% water for 30 minutes to overnight.
   3. Decant off the “fix” solution and rinse lightly with DI water
   4. Stain 30 minutes to overnight in Coomassie Brilliant Blue
   5. Decant off Coomassie Brilliant Blue and rinse with DI water
   6. Soak in DI water with trace amounts of ethanol to remove background staining. If gel is over stained, repeat step b and stain again for a shorter duration

2.5. Isotopically labeled ¹⁵N-AcpP

2.5a. Growth and Protein Expression:

1. Using pet29a C-terminal His₆-tagged AcpP that has been transformed into BL21 cells, prepare a 5mL starter culture in Luria-Bertani (LB) media with 5ul of 100 mg/mL ampicillin. Allow to incubate with rotation overnight at 37°C for a minimum of 12–18 hours.

2. Centrifuge the overnight culture at 12,000 rpm and 4C for 2 minutes to get a cell pellet. Discard the supernatant.

3. Resuspend the cell pellet it 1mL of sterile ultrapure water. Centrifuge under the same conditions in step 2. Discard supernatant. Repeat once more to ensure all LB has been removed. Resuspend a final time in 1mL of ultra-pure water **Washing the cells in ultra-pure water will induce the cytolysis of some cells. This can result in longer growth times. This cytolysis can be reduced by replacing ultra-pure water with M9 minimal media in this step**

4. Transfer 1L of sterile filtered M9 minimal media with 1g ¹⁵N-NH₄Cl to a clean, dry, autoclaved 2L baffled flask.

5. Add the resuspended starter culture and 1mL of 100mg/mL ampicillin to the baffled flask with M9 minimal media

6. Grow at 37°C with shaking of 150pms until OD₆₀₀ reaches 0.8

7. When OD₆₀₀ is reached, induce protein expression with 500uL of 1M IPTG for a final concentration of 0.5mM IPTG

8. Leave shaking at 150prm and 37°C overnight, for 12–18 hours

9. The following day, harvest the cell pellet with centrifugation for 45 minutes at 4°C and 2000 rpm. Discard the supernatant. **harvested pellets can be stored at ℒ20°C or used immediately**

2.5b. Protein Purification:

1. If cell pellet was previously frozen, thaw on ice

2. Resuspend cell pellet in lysis buffer

3. Sonicate the resuspended pellet for 6–8 minutes with intervals of 1s on and 4s off. Sonication should be performed over an ice bath and closely monitored to ensure the sample does not warm

4. Collect the lysate through centrifugation for 45 min at 12,000 RPM and 4°C. Transfer clarified lysate for protein purification. Discard pelleted membranes and insoluble materials

5. Batch-bind the lysate with Ni-NTA resin that has been equilibrated with lysis buffer. Use approximately 0.5mL of resin for 1L of growth. Allow to batch-bind for 30 minutes with spinning at 4°C

6. Prepare the following buffers and keep on ice.

   1. Wash 1: 50 mL of lysis buffer
   2. Wash 2: 50 mL of lysis buffer with 10mM imidazole (49.5mL of lysis buffer with 500uL of 1M imidazole added)
   3. Elution buffer: 50 mL of lysis buffer with 250mM imidazole (37.5mL lysis buffer with 12.5mL 1M imidazole added)

7. Transfer lysate to an empty gravity flow column fitted with a stop cock. Collect the flow-through. The Ni-NTA will form the bed volume as the flow through passes through the column. Do not allow the bed to go dry and stop the flow just above the Ni-NTA resin. Keep on ice after collection.

8. Wash with 40mL of Wash 1 buffer. Collect wash and stop flow just above the Ni-NTA resin. Keep on ice after collection.

9. Wash with 40 mL of Wash 2 buffer. **Bradford reagent can be used here to denote when to move to the elution buffer. The volume of Wash 2 can be increased or decreased as needed. Collect 20uL of sample and add 180uL of Bradford reagent. When no color change is detected, this indicates the majority of contaminant proteins have been removed and purification can proceed to the next step**

10. Elute protein with Elution buffer in 5mL factions testing by Bradford reagent as explained in the previous step.

11. Verify the presence and purity of the protein by testing the flow-through, wash 1, wash 2 and elution fractions by 12% SDS-PAGE gel.

12. To remove imidazole, dialyze fractions containing ¹⁵N-AcpP in 2L of 50mM Tris, 150mM NaCl, 10% glycerol, 1mM DTT and pH 7.4 overnight at 4°C.

2.6. Chemoenzymatic modification of ¹⁵N-AcpP:

In vivo expression of AcpP produces the ACP in two forms, apo- and holo-. Previous research has developed chemical biology tool kits to elucidate ACP mediated interactions via the loading of substrate mimetics to form crypto-AcpP, or by loading fatty acids to form acyl-AcpP [2, 7–13, 17–20]. It is first necessary to produce a homogenous sample of the carrier protein in the desired state, and then one of two methods can by applied for either cryptofication or acylation (Figure 2). Both methods are described below.

Figure 2:

AcpP is produced as a mixture of the inactive apo-ACP form and the active holo-ACP form. Homogenous holo samples are achieved using the phosphopantetheine transferase (Sfp) where homogenous apo samples are created with the phosphopantetheine hydrolase (AcpH). Synthesized phosphopantetheine mimetics can be loaded on to apo samples via a chemoenzymatic one-pot to produce non-native crypto-ACP where the native thioester bond has been replaced with an amide bond. The acyl-acyl carrier protein AasS can be used to attach free fatty acids to the holo-ACPs to produce acyl-ACPs that retain the native thioester bond. [7, 8, 15, 16]

2.6a. Holofication and acylation:

1. Prepare a uniformly holo-¹⁵N-AcpP sample

   1. Combine the following

      1. apo/holo-¹⁵N-AcpP mixture
      2. 0.005 equivalents of B. subtilis 4’-phosphopantetheinyl transferase (Sfp) **expression and purification as previously described** [17]
      3. 12.5 mM MgCl₂
      4. 1mM Coenzyme A
   2. Rotate overnight at 37°C
   3. Confirm complete holofication by Urea-PAGE analysis
2. Prepare acyl- ¹⁵N-AcpP

   1. Combine the following

      1. holo-¹⁵N-AcpP prepared in the previous step
      2. 0.005 equivalents of V. harveyi Acyl-acyl carrier protein synthetase (AasS) **expression and purification as previously reported** [7]
      3. 12.5 mM MgCl₂
      4. 8 mM ATP
      5. 0.5 mM TCEP
      6. 1.5 mM fatty acid
   2. Rotate overnight at 37°C
   3. Confirm complete acylation by Urea-PAGE analysis

2.6b. Apofication and cryptofication:

1. Prepare a uniformly apo-¹⁵N-AcpP sample

   1. Combine the following

      1. apo/holo-¹⁵N-AcpP mixture
      2. 0.1 equivalents of P. aeruginosa AcpH **expressed and purified as previously reported** [18]
      3. 12.5 mM MgCl₂
      4. 5 mM MnCl₂
      5. 5 mM TCEP
   2. Rotate overnight at 37°C
   3. Confirm complete apofication by Urea-PAGE analysis
2. Prepare crypto- ¹⁵N-AcpP

   1. Combine the following

      1. apo-¹⁵N-AcpP prepared in the previous step
      2. 0.1 equivalents each of E. coli CoaA, CoaD, and CoaE **expressed and purified as previously described** [19]
      3. 0.1 equivalents of B. subtilis 4’-phosphopantetheinyl transferase (Sfp) **expression and purification as previously described** [17]
      4. 12.5 mM MgCl₂
      5. 8 mM ATP
      6. 0.2% Triton-X
      7. 0.5 mM TCEP
      8. 1.5 mM substrate mimetic probe
   2. Rotate overnight at 37°C
   3. Confirm complete loading by Urea-PAGE analysis

2.8. Partner Proteins

Partner proteins should be expressed and purified as previously reported with the incorporation of the following modifications as necessary:

For NMR Titrations experiments using 2.6a, the active site of the partner protein may need to be mutated to prevent unwanted substrate modification. Further discussion can be found as previously reported. [1,7,16, 20]

2.7. FPLC Purification and NMR Buffers

After preparing the substrate loaded ¹⁵N-AcpP it will need to be isolated and transfered into a NMR suitable buffer solution.

2.7a. ¹⁵N-AcpP:

1. Separate by size exclusion chromatography. As AcpP is an 8.64 kDa protein, the Superdex S75 provides sufficient separation.

2. Buffer composition: 50mM phosphate, 0.5mM TCEP, 0.1% azide, pH 7.4

2.7b. Partner Protein

It may be necessary to test different buffer conditions for the chosen partner protein. The following should be taken into consideration when preparing the NMR buffer:

1. Stability: Titration experiments can require the sample to stay at experiment conditions for several hours to several days. The protein must be stable in the chosen buffer, under experimental conditions. Examples can be found as previously reported [7]

2. Salt: KCl and NaCl are often used to increase stability and solubility of proteins. However, the conductivity of these components interferes with the instrument’s signal. Phosphate 10mM-50mM is typically used to avoid conductivity interference with the signal to noise ratio

3. pH: Acidic pH is ideal (4–7) but the stability and longevity of the sample should be prioritized.

4. Reducing agents: 0.5–5mM DTT or TCEP can be used to prevent dimerization and unwanted activity due to the presence of free thiols

5. Microbial growth: Sodium azide <50uM can be used to prevent microbial growth in samples over the course of the experiments

**Note: Both the ¹⁵N-AcpP and the partner protein must be in the SAME buffer. The given ¹⁵N-AcpP buffer will need to be tested for compatibility with the partner protein. If adjustments are necessary to maintain protein stability, the determined buffer conditions should be uniformly applied to both ¹⁵N-AcpP and the partner protein.**

3. Materials and Methods for NMR Analysis

3.1. NMR Sample Preparation

The ¹⁵N-AcpP and partner protein samples must be prepared, then transferred to a high-quality NMR tube with a frequency rating of at least 500MHz, though 600MHz is preferable.

3.1a. Sample requirements

1. ¹⁵N-AcpP concentration: No less than 100uM

2. Partner protein concentrations: molar equivalents of 0.5, 1, 1.5, 2 with respect to ¹⁵N-AcpP

3. 10% D₂O in each sample

4. The total sample volume should be 500uL for a 5mm tube.

3.2. NMR Spectra Collection

Spectra should be collected on a 600 or 800MHz instrument such as the Bruker Advance spectrometer. The following protocol contains commands and parameters for use with TopSpin. TopSpin commands are shown in bold. All NMR experiments should be prepared, parameterized, and collected by, or under the supervision of, a person who has the appropriate expertise in NMR spectroscopy and the facilities being used.

1. Set the temperature to 37°C by using the command edta.

2. Load the sample of onto the instrument. **The “zero-point” sample of homogenous ¹H-¹⁵N-AcpP should be collected first. Experiments consisting of ¹H-¹⁵N-AcpP with increasing concentrations of partner protein will be conducted subsequently.**

3. Use the lock command to lock the sample on the deuterium signal frequency.

4. Tuning and matching the signal frequency are important for NMR experiments. If the instrument and software allow automated tuning, use the command atma for tuning and matching of the H, N, and C channels as necessary for the experiment. If manual tuning is required, display the wobble curve with the wobb command.

5. Shimming is a process that optimizes field homogeneity. The TopSpin command for shimming is topshim, though it can vary with instruments so it is important to verify the appropriate command.

6. Create a dataset to store output files with spectra data and specify parameters

7. Run a proton 1D experiment with water suppression to check if protein is folded correctly

   1. Parameters that need to be reviewed. The command ased will display the summary of acquisition parameters. The command eda will show all parameters.

      1. Recycle delay: 1.3s to 1.5s is recommended
      2. 90-degree pulse sequence calibrated to properly acquire signal from sample: getprosol command will populate the values. To find the accurate 90-degree pulse, use the pulsecal command
      3. Set O1 to water frequency
      4. Number of scans: You will need to specify both dummy scans (DS) and number of scans (NS). 16 scans should be sufficient, though number of scans will depend on the concentration of the sample. More dilute samples will need more scans for better sensitivity.
8. After 1D experiment is complete, create a new dataset to prepare the 2D HSQC

   1. Parameters that need to be reviewed. The command ased will display the summary of acquisition parameters. The command eda will show all parameters.

      1. Recycle delay: 1.3s to 1.5s is recommended
      2. 90-degree pulse sequence calibrated to properly acquire signal from sample: getprosol command will populate the values. To find the accurate 90-degree pulse, use the pulsecal command
      3. Set O1 to water frequency
      4. Data points: 2048 data points for the ¹H dimension, and 128 data points for ¹⁵N and ¹³C dimensions are recommended. More data points can be used to achieve better resolution.
      5. Number of scans: You will need to specify both dummy scans (DS) and number of scans (NS). 16 scans for the zero-point experiment is recommended and 24 scans is recommended for the samples with both the carrier protein and the partner protein. Number of scans will depend on the concentration of the sample. More dilute samples will need more scans for better sensitivity.
      6. Set your spectral widths (sw) and adjust the offset frequency as necessary for both proton and either N or C

9. The command rga sets the receiver gain

10. The command zg starts the data acquisition

11. The command xfb will process the 2D spectrum after acquisition.

12. Further processing can be performed though NMRPipe. [21]

3.3. NMR Data Analysis

3.3a. Chemical Shift Perturbations

Analysis of multiple HSQC spectra can be achieved using additional software designed to visualize and process different types of NMR data. As shown in Figure 3, by looking at chemical shift perturbations (CSP), information on individual residues, particularly residue specific interactions (Fig 3b and 3c) and key surface areas (Fig 3d), can be obtained. For titration experiments, there is a generalized workflow that ultimately calculates the chemical shift perturbations (CSP) per residue using the following formula:

$$\Delta {\delta }_{\mathit{\text{NH}}}=\sqrt{\Delta \delta H^2+{\left(\frac{\Delta \delta N}{5}\right)}^2}$$

Where $\Delta {\delta }_{\mathit{\text{NH}}}$ is the chemical shift for the NH bonds in the protein, while $\Delta \delta H$ and $\Delta \delta N$ represent the change in chemical shift for H and N nuclei, respectively. Backbone assignments, or peak lists, of the labeled protein are required for the proper assignment of residues to chemical shifts in the spectra. Commonly obtained through triple resonance assignments beyond the scope of this paper, the peak list can be downloaded from an online repository such as the Biological Magnetic Resonance Bank (BMRB). [22]

1. Obtain all necessary files pertaining to the HSQC spectra and convert them to an appropriate format for processing. Topspin saves the spectra in a particular format which can be converted to more accessible versions using a given NMR software. Popular software such as SPARKY, POKY (a SPARKY extension), or CCPN can be used for these purposes. [22–25]

2. Using either of the programs mentioned previously, load and visualize the zero-point spectra, modify contour as needed to obtain a clean view that only shows the NH signals and reduces noise.

3. Download the list of peaks from BMRB and map them to the spectra. Given slight differences in the buffers and other conditions, the peaks may appear slightly shifted from where they are supposed to be. Regardless, the overall fingerprint should appear the same. Manually adjust the peaks to center the signals observed that match the backbone.

4. NMR analysis software may be equipped with built-in functions that center the peaks using mathematical algorithms, this can help clear out bias once the peaks have been aligned with their corresponding chemical shifts.

5. Once all the peaks have been assigned to their corresponding residues, save the new peak list that contains the coordinates in the H and N dimensions for all the residues in the protein for the zero point.

6. Proceed to overlay the saturated spectra over the zero-point spectra and repeat the same steps for obtaining the coordinates that correspond to the saturated peaks. You may benefit by assigning different colors to each spectrum and go over each peak sequentially.

7. Save the saturated point spectra peak list as a separate file, and using excel or other numerical processing software, proceed to calculate the CSP for each residue in the protein.

**Note: Due to their lack of backbone NH bonds, prolines do not show up in the spectra. Histidine tags can also potentially interfere with HSQC spectra. Additionally, signal loss can be observed for some residues during data acquisition which may cause their peaks to be invisible.**

Figure 3:

Example figure of titration data of AcpP with E. coli LibB. A) The ¹H-¹⁵N-HSQC spectra were overlayed and individual peaks were highlighted. B) The CSP of each residue in the titration showing key residue specific interactions. C) Focus of the conserved Ser of AcpP D) CSP of AcpP colored by magnitude to show key surface areas. [10]

3.3b. Line Shape Analysis

NMR titration and binding experiments offer the ability to study structural and dynamical biomolecular interactions. Line shape analysis can be applied to two-dimensional experiments as unique chemical shifts are associated with each dimension of the experiment.[26] Line shape analysis allows the extrapolation of kinetic data such as exchange processes, k_ex, the dissociation constant, k_d, as well as the k_on and k_off rates.[26] This kinetic information can be determined for specific residues providing valuable insight into regulatory PPIs. Programs such as TITAN 2D have been designed for these purposes and can be used with the data from the acquired spectra. [7, 26, 27]

3.3c. Computational Analysis and Applications

Disrupting regulatory protein-protein interactions has become an enticing new target for rational drug discovery and design. High-throughput screening (HTS) is commonly employed in the pharmaceutical industry to screen large libraries of possible compounds, though its shortfalls include a high false-positive rate. This high failure rate combined with the complexity, time constraints, and high cost of drug discovery has contributed to an overall decline in the number of identified “possible” compounds successfully making it to market. [28] The development of methods, such as those described here, have unlocked valuable information that had previously remained elusive. NMR titration experiments on ¹⁵N labeled carrier proteins provide detailed structural information about the backbone NH bonds that are perturbed upon binding to partner proteins. [12] In addition to binding affinities, NMR titration experiments provide valuable insight into allosteric interactions, interfacial interactions, and confirmational changes that occur upon binding and complexation.[29] Each of these interactions offer new potential druggable sites. The quantitative amide proton CSP information provided though NMR titration experiments can be exploited to refine and improve docking simulations. By incorporating experimental restraints, an increased sampling of native-like confirmations can be generated, which in turn reduces the failure of scoring functions that rely on near-native conformation sampling. The augmentation of scoring/docking function with experimental restraints provides additional improvements in predictive power by overcoming deficiencies in current scoring schemes.[30] Unlike HTS, virtual screening (VS) and computer aided drug discovery (CADD) allow many molecules to be screened in a short amount of time though deficiencies in scoring functions still present limitations. Structure based virtual screening (SBVS) incorporates experimental and structural information allowing more refined screens with more optimized scoring functions.[31] Lead compounds or fragments identified through in silico methods that have been optimized with experimental restraints can then be tested in target-based and ligand-based NMR screens through additional NMR titration experiments.[31]