The overall goal of the following experiment is to study the mechanical properties and folding states of fusion protein binding partners using a FM based single molecule force spectroscopy. This is achieved by covalently immobilizing protein binding partners on a glass slide and on an A FM cantilever to achieve well-defined pulling geometry as a second step. The glass surface is indented with the cantilever, which allows receptor ligand binding to be established.
Next, the cantilever is retracted at a constant velocity in order to unfold individual protein domains and associate the receptor ligand complex results are obtained that reveal tertiary structural elements that act as energy barriers along the unfolding pathway, which are revealed in polymer elasticity models. Further kinetic and energetic information about the binding partners can be obtained by fitting kinetic models to the dynamic force spectrum. The overall goal of this procedure is to study the unfolding pathways of proteins under force.
This method can provide insight into the mechanical stability of proteins. It can be applied to any protein receptor ligand system that can be engineered with an accessible file group. Researchers using this method for the first time I struggle because timing is crucial and handling the cantilevers can be difficult.
This method can help answer key questions in the biophysics field, such as how a multi-domain protein unfolds To begin place. Two by 24 millimeter diameter 0.5 millimeter thick glass cover slips in A-P-T-F-E holder then sonicate. The cover slips in a 50 to 50 mixture of ethanol and ultrapure water for 15 minutes.
When sonication is complete, rinse the cover slips with ultrapure water and etch them in a piranha solution for 30 minutes. After 30 minutes, remove the cover slips and rinse them with ultrapure water and dry them under a gentle stream of nitrogen. Next submerge The cover slips in a 45 to five to one mixture of ethanol ultrapure water and three amino propyl dimethyl ahoy.
Place the submerge samples on a shaker at room temperature for 60 minutes at approximately 50 rotations per minute. Then dip the cover slip sequentially in 100%ethanol, followed by ultrapure water two times each. Again, dry the cover slips under a gentle stream of nitrogen.
Once dry, bake the cover slips in an oven at 80 degrees Celsius for 30 minutes. Store the unused cover slips under Argonne for up to six weeks to perform the amino ization of silicone cantilever tips. First place four silicone cantilevers on a clean glass slide and treat them with UV ozone for 15 minutes.
Then submerge cantilevers for three minutes and a 50 to 50 solution of ethanol and three amino propyl dimethyl oxis Ilene, along with point 25%ultrapure water. Next, rinse the cantilevers sequentially for 60 seconds in bakers of toluene, ethanol and ultrapure water. Carefully dry the cantilevers on filter paper between RINs after the final rinse.
Place the cantilevers on a clean glass slide and bake them at 80 degrees Celsius for 30 minutes. Prepare two Ts of TEP diss sulfide reducing beads by first cutting the tip off of a 200 microliter pipette tip to widen its diameter and then pipetting 100 microliter of the stock bead solution into each 1.5 milliliter micro tube. Next, add one milliliter of TBS to the TEP bead slurry OTTs, and rinse the beads by briefly vortexing the micro tube.
Then pellet the beads by centrifuging the samples at 850 Gs for three minutes. Carefully remove and discard the supernatant with a micro pipette and rinse the beads twice more with one milliliter of TBS. Centrifuging between rinses is after the final rinse, add a concentrated protein solution at three milligrams per milliliter to the TEP beads in a one to two ratio and gently mix the slurry by stirring with a micro pipet tip to avoid introducing air bubbles.
Then place the protein TEP bead slurry mixture on a rotator for 2.5 hours. Soak amino eyes cantilevers and cover slips in sodium bate buffer at a pH of 8.5 for 45 minutes to de protonate the primary amine groups on the surface. Then warm N-H-S-P-E-G malam powder to room temperature and prepare 210 microliters of it at 25 micromolar by vortexing it in sodium boy eight buffer.
The solution should be used as quickly as possible due to the extremely short half-life of NHS. At pH 8.5, quickly deposit 30 microliter droplets of N-H-S-P-E-G malam solution onto a Petri dish and 90 microliters onto every other glass cover slip. Carefully place the cantilevers inside the 30 microliter droplets.
Then flip the cover slip without the droplet onto one with N-H-S-P-E-G malam solution to form a sandwich with the N-H-S-P-E-G MALAM solution. In the middle, incubate the cantilevers and cover slips with the N-H-S-P-E-G MALAM solution at room temperature for one hour. Centrifuge the TEP bead and reduced protein solution at 100 GS for one minute and collect the snat.
Next, dilute the protein solution with TBS. Aim for a protein concentration during surface conjugation in a range of 0.5 to two milligrams per milliliter. Then set the reduced and diluted protein solutions aside for a few minutes while rinsing cantilevers and cover slips.
Rinse the cover slips in three sequential beakers of ultrapure water. Remove residual liquid from the cover slips by carefully touching the edges to filter paper. Then mount the cover slips in an appropriate sample holder that is compatible with the A FM instrument and dry residual liquid under a gentle stream of nitrogen.
Apply 20 microliters of the diluted protein solution as a droplet onto the center of the cover slips. Then rinse the cantilevers in three sequential beakers of ultrapure water, followed by 30 microliter droplets of the second diluted protein solution. Incubate pegylated cover slips and cantilevers with respective diluted protein solutions at room temperature for one to two hours in a humidity chamber.
Next, use a pipette to rinse cover slips at least 10 times with one milliliter of TBS and then store them in TBS until the measurement. Also rinse the cantilevers using three sequential beakers containing TBS to remove unbound proteins and store them in TBS as well. Mount the functionalized, cantilever and glass slides in an A FM model that is suitable for measuring in liquids and supports an accessible speed range on the Z pieto of approximately 200 to 5, 000 nanometers per second.
When mounting the can minimize exposure to air, ensure that the surfaces stay covered with buffer during the entire process upon correct adjustment of the laser beam. Let the system equilibrate for at least 30 minutes to reduce any drift effects and readjust if necessary. Then begin taking measurements upon successful binding of the cohesion.
Docker in pair the recorded force distance traces shown here. Exhibit a characteristic peak pattern. Every peak in the trace represents the unfolding of one protein subdomain with the last peak corresponding to the dissociation of the receptor ligand complex.
The recorded force distance traces were then transformed to force contour length space and assembled in a barrier position histogram. The data show a contour length increment of approximately 89 nanometers, which can be unambiguously assigned to the unfolding of the xin domain to probe the energy landscape of the cohesive docker and interaction. A total of 186 data traces were obtained at four different pooling speeds.
The resulting dynamic force spectrum shown as large blue circles represents the most probable rupture forces at a given loading rate. These values were then fitted to the Bell Evans model, the fit yielded values for Koff and delta X, which were found to in good agreement with previously published results. Once mastered, the experimental part of this technique can be done in eight to nine hours.
Following this procedure site-specific mutagenesis can be performed in a follow-up experiment to determine amino acids crucial for binding After its development. This technique paved the way for researchers in the field of single molecule biophysics to explore protein folding landscapes. We use it to study the mechanical properties of cellulose SOEs, which are among the most mechanically stable protein systems known.
After watching this video, you should have a good understanding of how to perform single molecule force spectroscopy on protein receptor ligand pairs.
