Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy (Conpokal) on Live Cells

Conpokal combines confocal microscopy with atomic force microscopy using a probe to poke the sample surface. Although both techniques are effective individually, Conpokal facilitates fluorescence co-localization with mechanical characterization. The main advantage of the Conpokal technique, is the near simultaneous dual-microscopy.

Confocal investigates cytoskeletal and other cellular processes before and after AFM probing, which delivers area specific mechanical properties. This technique is impactful within the mechanobiology field as for example, brain cells can be probed under physiological conditions to examine electrical impulses and force transaction. Before beginning the analysis, select an appropriate AFM cantilever for the desired data collection and use gloves, the AFM chip mounting stage, tweezers, and a small screwdriver to mount the chip into the glass block.

Carefully place the AFM chip onto the center of the glass block. The cantilever plus a very small portion of the AFM chip should be in the visible, non-opaque portion of the glass block. Use the screwdriver to tighten the screw until the chip is snug against the glass block, and use a lens to check that the chip is oriented correctly.

When the chip has been properly oriented, place the glass block into the AFM head in the proper orientation and lock the glass block into place. After locating the bottom of the calibration dish by bright-field microscopy, use the AFM system Z stepper motor control panel to move the AFM cantilever 2000 microns above the sample. Using bright field illumination and the Z stepper motor control panel, slowly lower the AFM chip to the bottom of the glass dish in steps of 100 to 200 microns to avoid crashing the AFM tip into the Petri dish.

Locate the manual micrometers that control X Y or in plain motion of the AFM head on the instrument platform. As the shadow from the AFM cantilever becomes darker, and the shape becomes sharper, use the manual micrometers to correct the AFM head and adjust the position of the AFM cantilever within the field of view. At this time, the lookup tables may need to be adjusted.

Watch for a shadow to appear in the microscope software view, which indicates that the AFM tip is getting closer to the bottom of the dish. Continue using small steps to lower the AFM tip until the tip is mostly in focus. When the laser is in position, use the laser alignment dials to position the laser on the backside of where the AFM tip is located on the cantilever.

The laser alignment panel should display a sum signal greater than zero volts. Move the laser, a small distance in all of the directions on the AFM cantilever, until the maximum sum signal is achieved while remaining at the AFM tip. Once the laser position is set, use the manually controlled deflection dials to zero the vertical and lateral deflection.

Use the vertical and lateral deflection knobs to align the detector so that the target is centered and there is no vertical or lateral deflection observed in the laser alignment panel. Open the Calibration window, and enter all of the experiments specific information. Replace the laser light filter.

Before calibrating, turn off the confocal microscope light source and close the AFM enclosure to dampen any potential noise coming from the room light or vibrations. Press the Calibration button to automatically allow the system to calibrate the tip. When the calibration is complete, the stiffness of the cantilever and its sensitivity will be displayed in the Calibration panel.

Then use the Automated Approach Command button to lower the tip to the bottom of the sample dish and set the size of the skin area, the resolution, the set point, the Z length, and the pixel time, before pressing the play button to begin scanning. For confocal microscope imaging in the microscope software, enable the confocal capabilities and select the laser lines appropriate for the dyes that were used to stain the samples. Select one or multiple laser lines to excite and image those features in the sample and set the gain to a value that optimizes the sample fluorescence but limits the amount of noise.

Adjust the laser power to avoid saturated pixels while maximizing the dynamic range and set the pinhole size to one area unit to maximize the resolution for the optical sectioning. If needed, adjust the values based on the sample brightness. To set the pixel dwell time, begin with about a two microsecond dwell time, and adjust to reflect the sample brightness as needed.

To select the pixel size for the selected objective, let the instrument calculate the pixel size via the Nyquist option button and the selected number of pixels in the image. Next, select the Scan option and begin the data collection. Use the focus knob to locate the focal plane representing the samples features in its clearest form, the Scan button to begin data collection, and the Capture button to capture a two dimensional image.

Using the panel which controls the pixel size via the Nyquist option, reduce the area size of the scan to only envelop a single cell. Activate the collection tool and using a bottom to top option, with only the laser line that illuminates a feature in the sample most clearly, set the start and finish planes for the volume to be measured using the suggested spacing between the planes. Name the file and save it to the associated folder.

Alternatively, if our priori knowledge of the samples thickness exists, select the brightest, sharpest, middle plane by adjusting the focus to define the volume and set the top to have the sample thickness above the middle plane, and the bottom to have the thickness below the middle plane, then select the appropriate step size. Run the acquisition by pressing the Run Now button. When the acquisition finishes, save the file to the appropriate folder.

At the end of the experiment, wear gloves while removing the glass block and place it into the mounting station. Use tweezers with rubber grips to carefully remove the AFM chip and place the used tip back into the location of the storage box oriented off access to denote that it has been used. For future reference, note which tip was used in the experiment.

Here, a representative AFM scan over a living HEK cell is shown. The height for this particular HEK cell was around 10 microns as demonstrated by the line scan. Here, an example of a poor scan due to an improper AFM tip choice can be observed.

In this image, black pixels appeared at the apex of the cell indicating that the AFM PA zone was out of range due to a large cell height. The end of the AFM cantilever also appears in the image because of the tip offset combined with an insufficient tip height compared to the cell height. These artifacts in the AFM image, indicate that a different AFM tip should have been chosen to image the cell.

Three color confocal imaging can be performed. For example, to visualize the cell nucleus, micro tubules, and lipophilic membrane. Analysis of the tip indentations in combination with the nano mechanical model, allows the generation of a modulus map of the surface.

Here, a corresponding 3D projection of the laser confocal Z stack is shown. AFM scans and measured modulus maps can also be acquired for microbes, as observed in these representative analysis of streptococcus mutans bacterium. As demonstrated, a better resolution can be achieved at this scale with AFM, than with traditional confocal microscopy.

Adhesive force mapping is a technique performed through conpokal to visualize molecular interactions. For example, to study the modulation of cell surface molecules to observe binding kinetics. Conpokal ushers in a new pathway for exploring structure function relationships in medical microbiology.

For example, physical and chemical events and the peptidoglycan layer, can be linked to antibiotic resistance.