Protein-Protein interactions are important in virtually all biological process. Understanding of these reactions will aid in designing novel and effective drugs/. ACE is used as a high screening to establish and define the binding interaction between a therapeutic target protein and natural or chemical molecules. This technique is also used in the measuring the binding constants, estimation of kinetic rate constants and the stoichiometry of protein-protein interaction.
- 1 Principle
- 2 Materials
- 3 Methods
- 4 NMR SPECTROSCOPY
- 5 NMR Spectroscopy
- 6 NMR Chemical Shift Titration
- 7 Studying Protein–Protein Interactions via Blot Overlay or Far Western Blot
- 8 Methods
ACE is based on the electrophoretic separation of analytes, under soluble conditions in which the analyte carries a net charge. With proteins, CE conditions are established to a pH using buffers. This pH is different from proteins isoelectric point to prevent precipitation of pH. The rate at which the transfer occurs is target specific, generally, it is dependent on the ratio of charge to mass. Different proteins or isoforms migrate with an individual velocity. Detection is done with the photomultiplier tube. A target CE profile is created. This technique has high resolution. This fact is contributed to high field strengths and short run-time. The subnanomolar concentration of proteins can be calculated using laser-induced fluorescence detection. If a binding has occurred it alters the rate of migration of the protein, which will result in a different protein profile.
ACE requires a short analysis time. It requires only minutes. ACE is automatable for high-throughput screening. It can be used to find the relative strength of binding. It can distinguish the targeted protein from background fluorescence. This problem occurs in the case of natural crude extracts.
- Capillary Electrophoresis Unit.
- Borosilicate capillary: 50 μm, 75 μm, 100 μm diameters.
- Surfactants: Brij-35, Triton X-100, and CHAPS.
- Fluorescein isothiocyanate (FITC).
- Protein labeling reagents (Molecular Probes).
- Vascular endothelial growth factor (VEGF-165).
- Vascular endothelial growth factor receptor (VEGF-r1).
- Stock buffer: 20 mM Tris-HCl, pH 8.0, 5 mM DTT, 0.1% CHAPS, 150 mM NaCl.
- Sample buffer: 20 mM Tris-HCl, pH 8.3, 1 mg/mL BSA.
- Running buffer: 50 mM Tris-HCl, pH 8.3, 1 mg/mL BSA.
- Micro-spin columns.
- Assay reagent (Bio-Rad).
General Instrument Set Up
CE was performed on a Capillary Electrophoresis instrument equipped with a diode array detector or LIF detection (488 nm excitation, 520 nm emission). The capillary is composed of fused silica with 50 μm, 75 μm, 100 μm diameters and a total length of 20 cm. The length of run from the inlet to the detectors in 12 cm. Capillaries were coated on the internal walls with a polymer to prevent protein absorption and reduce electro osmotic flow (EOF). Operating Voltage values ranges from 15-30 kV. Hydrodynamic injection of the sample was performed at a pressure of 0.5 psi for 10–20 s. The capillary temperature was maintained at 20°C. Data retrieval and analyses were performed by P/ACE MDQ Software
Assay for Growth Factor/Receptor Interaction
Growth factor–growth factor receptor interactions are the initial step of many signal transduction events the direct cell growth, differentiation and apoptosis. Vascular endothelial growth factor (VEGF) and its receptor interaction is studied in this example using ACE.
In this assay, VEGF-165 was labeled with FITC in 30 mM HEPES buffer using a dye to protein molar ratio of 10:1. Labeled target was separated from free dye using microsep spin columns. The concentration of the purified material was obtained by using the Bradford Assay with BSA as a standard. Equal molar amounts of VEGF and VEGFr1 (50 nM of each in stock buffer) were used in these experiments by dilution of each into Tris based sample buffer. The sample was kept at 4°C prior to injection onto the capillary. Crude natural extracts in 100% DMSO were added to the running buffer to the level of 1%; clean DMSO was used as a control.
Data from the CE Assay
Within the capillary, labeled VEGF migrates within a large segment of BSA that is provided with the target protein. When VEGF is labeled in a mixture containing excess BSA, then BSA will also be labeled. Thus, the unbound labeled VEGF is not visible in the CE electropherogram due to excess labeled BSA. Upon the addition of unlabeled VEGF-r1, the VEGF/VEGF-r1 complex shifts away from the labeled BSA and is visible as a separate peak.This slower migration is expected as the receptor is basic and its binding to VEGF would provide for a complex of slower mobility than the growth factor alone under these conditions.
Data Analyses from the CE Assay
In theory, the fluorescently labeled (FL)-VEGF/VEGF-r1 complex CE profile would be sensitive to hits interfering with complex formation and one might hypothesize that molecules that interfered with binding would manifest as a loss in signal of the complex peak. As the concentration of unlabeled VEGF is increased, it competes with the labeled VEGF for binding to the VEGF-r1, resulting in increasing amounts of unlabeled-VEGF/VEGF-r1 complex formation and a decrease in the amount of FL-VEGF/VEGF-r1. This results in an overall loss of labeled complex signal.These data may be interpreted to indicate that sample, which caused an overall loss of labeled complex signal, would be selected for further study (fractionation, testing of fractions for activity, and isolation of chemical entity.) In screening a large number of crude extracts, we found the primary hit rate using this format to be approx 1%.
- 15N-labeled Raf-CRD and unlabeled Ras.
- NMR buffer: 30 mM d11-Tris-d3-acetate (Isotec, Miamisburg, OH), pH 6.5, 100 mM NaCl, 10 μM ZnCl2, 1 mM d10-DTT, 10% D2O, 0.01% NaN3.
- Bruker AMX 500 NMR spectrometer or Varian Inova 600 MHz NMR spectrometer.
- 5.0 mm Wilmad standard NMR sample tube.
- FELIX version 97.2.
NMR Sample Preparation
The Raf-CRD has expressed as glutathione S-transferase (GST) fusion protein and purified by affinity chromatography. Uniformly 15N enriched Raf-CRD protein was obtained by growing Escherichia coli in a supplemented minimal media with 99.8% 15 NH4Cl as the sole nitrogen source. The purified protein was concentrated to 2 mL in 30 mM Tris acetate at pH 6.5, 75 mM Na2SO4, 10 μM ZnCl2, 1 mM DTT and was exchanged into NMR buffer 30 mM d11-Tris-d3-acetate at pH 6.5, 100 mM NaCl, 10 μM ZnCl2, 1 mM d10-DTT, 10% D2O, 0.01% NaN3] using a gel filtration column. All NMR samples had the final volume of 500 μL. The Raf- CRD NMR sample contained 0.28 mM uniformly 15N-enriched Raf-CRD alone or mixed with wild-type unlabeled Ras at concentrations ranging from 0.30 to 0.77 mM.
NMR experiments were recorded on a Bruker AMX 500 (500 MHz) spectrometer at 12°C or a Varian Inova 600 (600 MHz) spectrometer at 25°C. Two-dimensional 1H–15N heteronuclear single quantum coherence spectroscopy (HSQC) experiments were performed with pulse field gradient and water flip-back methods. Data were acquired with 1024 × 256 complex data points and a spectral width of 7042.25 Hz for the 1H dimension and 1000 Hz for the 15N dimension on the Bruker AMX 500. HSQC experiments conducted on the Varian Inova 600 spectrometer were acquired with 1024 × 128 complex data points and a spectral width of 8000 Hz for the 1H dimension and 1700 Hz for the 15N dimension. NMR data were processed and analyzed using the program FELIX.
NMR Chemical Shift Titration
Proton and 15N resonance assignments of the Raf-CRD were determined previously in this laboratory. Two-dimensional 1H–15N HSQC data were acquired on the Raf-CRD-Ras complexes at four different molar ratios: 1:2.75, 1:1.65, 1:1.1, and 1:0. The final concentration of 15N-Raf-CRD was 0.28 mM and unlabeled Ras ranged from 0.77 to 0.30 mM. Samples containing the highest molar ratio (1:2.75) of the Raf-CRD–Ras complex (0.28:0.77 mM) and Raf- CRD alone (0.28 mM) were prepared first. To prevent changes in sample volume and buffer composition, the other two complexes (molar ratio 1:1.65 and 1:1.1) were made by exchanging 200 μL of the two samples containing the highest molar ratio (1:2.75) and the sample alone (1:0) to achieve a final sample volume of 500 μL. A fluorescence-based binding assay was used to determine the binding affinity between the Raf-CRD and Ras. Owing to solubility limitations, a quantitative measure of the Kd could not accurately be determined. However, our estimated Kd of 300 μM indicated that the binding interaction between Raf-CRD and Ras was likely to be in fast exchange on the NMR time scale. During the titration, we expected to see only one peak that represents a weighted population of free and bound states. Given an estimated Kd of 300 μM for the Ras/Raf-CRD interaction, the NMR experiments were conducted under conditions ranging from approx 0 to 66% occupancy, and the full extent of chemical-shift changes (full saturation binding) will not be sampled. However, if our Kd is underestimated, our fractional occupancy will be less than 66%. Even though the chemical-shift changes will be smaller than would be anticipated for a titration that reaches saturation binding, these experiments, nevertheless, report chemical-shift changes that allow mapping of the binding interface.
To elucidate the residues of the Raf-CRD important for Ras binding, we evaluated 1H–15N HSQC NMR spectral changes associated with binding of Mapping Protein–Protein Interactions by NMR 83 Ras to the 15N-enriched Raf-CRD. The 1H–15N HSQC experiment will give rise to a spectrum of proton chemical shifts in one dimension and the chemical shifts of the covalently attached 15N nucleus in the second dimension, so only peaks corresponding to 15N-enriched Raf-CRD will be observed. Among the various HN resonances detected are the amide protons. Therefore, the HSQC peaks provide a site-specific probe for every residue in Raf-CRD with the exception of proline. NMR spectral changes in the Raf-CRD after the addition of Ras (molar ratio 1:2.75, as a superposition of a portion of 1H–15N HSQC spectra of uniformly 15N-labeled Raf-CRD in the presence (black) and absence (gray) of unlabeled Ras. Raf-CRD residues that show distinguished changes in 1H and 15N chemical shift and/or peak intensity are labeled.
Most of the chemical-shift differences observed between the free Raf-CRD and the complex are localized to Raf-CRD residues 144, 145, 148–150, 158–164, and 174–176. The largest 1H chemical-shift changes observed showed slightly smaller chemical-shift changes. The average change of the 1H chemical shifts for the rest of the peaks was less than the line width, indicating that the global structure of Raf-CRD is unchanged due to the association with Ras. The same trend was observed in the 15N chemical shift changes. Uniformly reduced peak intensities (approx 55%) were also observed for a number of peaks, which likely reflects additional relaxation mechanisms due to interaction with the Ras protein. The largest variation in intensity changes for the amide peaks was found for residues 141, 145, 148–150, 158, and 170. Clearly, residues 148–150 and 158–164 at the top of the molecule are identified as part of the binding interface. Consistent with these observations, an L149T/F151Q Raf-CRD mutant exhibits reduced binding affinity to Ras in vitro. Moreover, the L149T/F151Q mutation when placed in the context of full-length Raf showed impaired Ras-mediated Raf activation in vivo (4).
Studying Protein–Protein Interactions via Blot Overlay or Far Western Blot
During preparation for SDS-PAGE, proteins are typically reduced and denatured via treatment with Laemmli sample buffer. Because many protein– protein interactions rely on aspects of secondary and tertiary protein structure that are disrupted under reducing and denaturing conditions, it might seem likely that few if any protein–protein interactions could survive treatment with SDS-PAGE sample buffer. Nonetheless, it is well known that many types of protein–protein interaction do in fact still occur even after one of the partners has been reduced, denatured, run on SDS-PAGE, and Western blotted. Blot overlays are a standard and very useful method for studying interactions between proteins.
In principle, a blot overlay is similar to a Western blot. For both procedures, samples are run on SDS-PAGE gels, transferred to nitrocellulose or PVDF, and then overlaid with a soluble protein that may bind to one or more immobilized proteins on the blot. In the case of a Western blot, the overlaid protein is the antibody. In the case of a blot overlay, the overlaid protein is a probe of interest, often a fusion protein that is easy to detect. The overlaid probe can be detected either via incubation with an antibody (this method is often referred to as a “Far Western blot”), via incubation with streptavidin (if the probe is biotinylated), or via autoradiography, if the overlaid probe is radiolabeled with 32P.
- SDS-PAGE mini-gel apparatus (Invitrogen).
- SDS-PAGE 4–20% mini gels (Invitrogen).
- Western blot transfer apparatus (Invitrogen).
- Power supply (BioRad).
- Nitrocellulose (Invitrogen).
- SDS-PAGE pre stained molecular weight markers (BioRad).
- SDS-PAGE sample buffer: 20 mM Tris-HCl, pH 7.4, 2% SDS, 2% β-mercaptoethanol, 5% glycerol, 1 mg/mL bromophenol blue.
- SDS-PAGE running buffer: 25 mM Tris-HCl, pH 7.4, 200 mM glycine, 0.1% SDS.
- SDS-PAGE transfer buffer: 10 mM Tris-HCl, pH 7.4, 100 mM glycine, 20% methanol.
- Purified hexahistidine-tagged fusion proteins.
- Purified GST-tagged fusion proteins.
- Anti-GST monoclonal antibody (Santa Cruz Biotechnology, cat. no. sc-138).
- Goat anti-mouse HRP-coupled secondary antibody.
- Blocking buffer: 2% nonfat powdered milk, 0.1% Tween-20 in phosphate-buffered saline, pH 7.4.
- Enhanced chemiluminescence kit.
- Blot trays.
- Rocking platform.
- Autoradiography cassette.
- Clear plastic sheet.
3.1. SDS-PAGE and Blotting
The purpose of this step is to immobilize the samples of interest on nitrocellulose or an equivalent matrix, such as PVDF. It is very important to keep the blot clean during the handling steps involved in the transfer procedure because contaminants can contribute to increased background problems later on during detection of the overlaid probe.
- Place gel in SDS-PAGE apparatus and fill the chamber with running buffer.
- Mix purified hexahistidine-tagged fusion proteins with SDS-PAGE sample buffer to a final concentration of approx 0.1 μg/μL of the fusion protein.
- Load 20 μL of the fusion protein (2 μg total) in each lane of the gel. If there are more lanes than samples, load 20 μL of sample buffer in the extra lanes.
- In at least one lane of the gel, load 20 μL of SDS-PAGE molecular weight markers.
- Run gel for approx 80 min at 150 V using the power supply.
- Stop gel, turn off the power supply, remove the gel from its protective casing, and place in transfer buffer.
- Place pre-cut nitrocellulose in transfer buffer to wet it.
- Put nitrocellulose and gel together in transfer apparatus, and transfer proteins from gel to nitrocellulose using a power supply for 80 min at 25 V.
During the overlay step, the probe is incubated with the blot and the unbound probe is then washed away. The potential success of the overlay depends heavily on the purity of the overlaid probe. GST and hexahistidine-tagged fusion proteins should be purified as extensively as possible. If the probe has many contaminants, this may contribute to increasing the background during the detection step, making visualization of the specifically bound probe more difficult.
- Block blot in blocking buffer for at least 30 min.
- Add GST fusion proteins to a concentration of 25 nM in 10 mL blocking buffer.
- Incubate GST fusion proteins with blot for 1 h at room temperature while rocking slowly.
- Discard GST fusion protein solution and wash blot three times for 5 min each with 10 mL of blocking buffer while rocking the blot slowly.
- Add anti-GST antibody at 1:1000 dilution (approx 200 ng/mL final) to 10 mL blocking buffer.
- Incubate anti-GST antibody with blot for 1 h while rocking the blot slowly.
- Discard anti-GST antibody solution and wash blot three times for 5 min each with 10 mL of blocking buffer while rocking the blot slowly.
- Add goat anti-mouse HRP-coupled secondary antibody at 1:2000 dilution to 10 mL blocking buffer.
- Incubate secondary antibody with blot for 1 h while rocking the blot slowly.
- Discard secondary antibody solution and wash blot three times for 5 min each with 10 mL of blocking buffer while rocking the blot slowly.
- Wash blot one time for 5 min with phosphate buffered saline, pH 7.4.
3.3. Detection of Overlaid Proteins
The final step of the overlay is to detect the probe that is bound specifically to proteins immobilized on the blot. In viewing different exposures of the visualized probe, an effort should be made to obtain the best possible signal-to-noise ratio. Nonspecific background binding will increase linearly with time of exposure. Thus, shorter exposures may have more favorable signal-to-noise ratios.
- Incubate blot with enhanced chemiluminescence solution for 60 s.
- Remove excess ECL solution from blot and place blot in clear plastic sheet.
- Tape sheet into autoradiography cassette.
- Move to a darkroom and place one sheet of film into autoradiography cassette with the blot.
- Expose film for 5–2000 s, depending on the intensity of signal.
- Develop film in standard film developer.