The overall goal of this procedure is to achieve the highest possible resolution imaging in liquid, with a commercial AFM operated and amplitude-modulation, also known as tapping mode. This method helps pushing the limit of standard AFM operation in liquid, by using the best combination of parameters for high resolution. The main advantage of this technique is that it can be used with most commercial AFM's, and as such it doesn't require any specialist equipment.
The method presented here is aimed at scientists and technician who already have some basic knowledge of AFM, but would like to get more out of the technique. This method is not aimed at any particular type of sample, and can be broadly applied to samples from physics, biology, chemistry, materials, and service sciences. Generally, individuals new to this technique will struggle, because it requires some patience to find the best parameters for a given sample.
Bath sonicate the instruments, and the disk supporting the substrate in ultra pure water. Followed by isopropanol, and again ultra pure water, each for 10 minutes. When aiming for high resolution, any contamination can have detrimental consequences.
Wear gloves at all times, and ensure that any surfaces or instruments that come in contact with the sample, cantilever, or AFM cell, are thoroughly cleaned. After sonication, dry each of the instruments and the sample disk under a flow of nitrogen. Use a steel disk as the support for mica, to image single absorbed metal ions.
Physically clean the surfaces that can't be cleaned by sonication by wiping them with single ply low lint tissues soaked in ultra pure water, isopropanol, and ultra pure water sequentially. Allow the surface to dry in air for up to 30 minutes. Next, prepare a small amount of epoxy glue by thoroughly mixing the reagents, and place about 10 microliters of the epoxy on the clean steel sample disk.
Place the mica substrate on the epoxy, and affix it to the steel disk by applying pressure on the substrate. Allow the epoxy to cure for several hours at an elevated temperature according to the manufacturer specifications. Then, firmly press a 2.5 centimeter wide piece of adhesive tape onto the substrate, so that the entire face is covered, and smoothly peel off the top layer.
Repeat this process two to three times until the mica is mirror smooth to the eye. Immerse the cantilever chip in a bath of isapropanol followed by ultra pure water each for 60 minutes. Then, expose the tip to UV light for up to five minutes in order to favor the formation of stable hydration sites.
Longer over-exposure times can damage the tip, or increase its radius of curvature. Insert the cantilever into the AFM's cantilever holder, and pipette 25 microliters of the imaging liquid onto the cantilever and tip to pre wet the surface. This will reduce the appearance of air bubbles when approaching the sample.
Melt the sample disk and substrate onto the sample stage and add a droplet of the imaging liquid to the sample. Then connect the cantilever holder to the AFM. Bring the cantilever and sample into close proximity so as to form a capillary bridge between the fluids on the cantilever tip and those on the sample.
Use the AFM software to align the measuring laser close to the tip end of the cantilever. Next, find the residence frequency of the cantilever from the main peak in its thermal spectrum. If the deflection of the cantilever is calibrated, fitting the residence peak with a simple harmonic oscillator model yields the spring constant of the cantilever.
Then, tune the cantilever by finding its amplitude response when externally driven over a range of frequencies close to the resonance frequency identified in the thermal spectrum. Adjust the driving amplitude so that the free oscillation amplitude is approximately five nanometers. This typically corresponds with 0.2-0.8 volts on most AFM's.
Then, adjust the amplitude set point to about 80%of the free amplitude. Next, set the feedback gains relatively high. After ensuring that no instability or ringing occurs, set the initial scan rate to about one hertz, and the scan size to 10 nanometers.
Initiate the tip's approach to the surface using the AFM control software. Assess wether the tip has reached the surface without starting to image by slightly changing the set point value. If the tip is at the surface, the effect on the extension of the ZPA zone should be negligible.
Once the tip has reached the surface, retract the ZPA zone and retune the cantilever. The resonance frequency will likely have shifted to a lower value due to tip sample interactions. Now, change the set point to about 80%of the newly tuned free amplitude, and engage the cantilever to conduct a 10 by 10 nanometer squared scan of the surface in amplitude modulation mode to verify that the imaging parameters are suitable.
Check that the trace and retrace profiles superimpose. If not, further reduce the set point, and try increasing the gains. If the image becomes noisy, lower the gains.
Repeat the operation with a large one to five micrometer squared region of the sample, provided this is possible. On soft or biological samples, this might result in contamination of the tip. Reduce the scan size to a value suitable for visualizing the features of interest.
This can be as low as 20 by 20 nanometers. Next, reduce the drive amplitude of the cantilever enough for the feedback loop to automatically retract the ZPA zone, enhance the tip from the surface. While the cantilever is away from the surface, adjust the drive amplitude so that the cantilever amplitude is one to two nanometers peak to peak.
Progressively reduce the set point in small steps, until the ZPA zone extends again towards the surface, and the original image is recovered. Keep the set point amplitude between 75%and 95%of the new free amplitude. Then, readjust the gains, since higher gains can be used at lower amplitudes without introducing significant noise.
Optimize the system to find the best combination of free amplitude, set point, and gain for high resolution. The optimum system conditions depend on the sample, the wetting properties of the liquid, and also the cantilever being used. For solvophilic interfaces, use cantilevers with a spring constant of 0.5 to two newtons per meter.
Using this technique, sub nanometer resolution images were obtained over a broad range of samples. The soft samples shown here include a lipid bilayer, purple membranes from halobacterium salinarum, a self assembled monolayer of amphophilic dimolecules, and aquaporin crystals from a bovine lens membrane. In each case, the features of interest are highlighted.
The small oscillation amplitudes and high set points minimize the force exerted by the tip on the sample, allowing for the fragile self assemblies of lipids in the bilayer, proteins in native biomembranes, and amphophilic molecules to be imaged in solution without damage. Harder crystalline materials, such as calcite, strontium titanite, silicon carbide, and single metal ions absorbed on a mica surface can be imaged using this approach, because in every case, it is the interfacial liquid that is effectively imaged, not the crystal itself. Once mastered, this technique provides molecular or atomic level resolution in liquid almost every time it's performed correctly.
When attempting this procedure, it's important to bare in mind that it's the interfacial liquid being imaged. This means using soft imaging conditions. Contamination of the tip, the sample, or the liquid is usually the main cause for failing to achieve high resolution.
If in any doubt, it is often a good idea to clean all the surfaces in contact with the liquid, and reuse imaging solutions. External noise is also detrimental to high resolution. A low vibration floor, away from a ventilation duct is better.
After watching this video, you should have a good understanding of how to optimize your imaging parameters to achieve high resolution AFM. Of course, as for any cutting-edge technique, it can take several trials to figure out how to best image a sample, so patience is key.
