In the DC mode, a continuous feedback mechanism moves the pipette up and down while scanning the sample to keep the pipette always in the proximity of the sample surface. Thus, a pipette-sample separation, typically equal to the pipette's inner radius, is maintained during imaging. This method suffers from poor sensitivity and resolution. Moreover, in principle, the probe does not come into contact with the surface. However, since the dependence of the ionic current on the distance between the tip and the sample is not very steep, the tip often contacts the surface of a rough sample during routine imaging. Therefore, the DC mode is now generally used only for sample with vertical protrusions below a few hundred nanometers.

In the AC mode, a lock-in amplifier is used to detect the changes in the AC current amplitude, which is then used to control the feedback circuitry to modulate the scanning tip distance from the sample surface. This mode is much more sensitive than the DC mode, and allows the probe to be operated only a few nanometers from the sample surface. Moreover, the AC ionic current can be recorded simultaneously during the scan. As described in a previous report (Pastre et al., 2001), it can provide additional information about the local ionic environment. However, it is impossible to scan highly convoluted surfaces using either the DC or the AC mode because the SICM probe only senses locally at the tip. In other words, in these two types of mode, the side of the probe may touch the sample before the tip has sensed the presence of a very bumpy surface. Therefore, like the DC mode, the AC mode is also only suitable for samples with a relative smooth surface.

The hopping mode is also called a backstep or ARS mode. In the hopping/ARS mode, ion current is recorded while the pipette is moved vertically and repeatedly approaches and retracts from the sample surface. This new mode type no longer uses continuous feedback and raster scanning pattern. The nanopipette approaches the surface to measure the height only at selected imaging points and rapidly retracts back to a safe distance before moving laterally onto the next imaging point. The pipette approaches until the current is reduced by a predefined threshold. Typically, the pipette is still at a distance of about one inner pipette radius from the surface, usually 25–50 nm. Therefore, the hopping mode is especially powerful for imaging samples with steep slopes and is capable of imaging surfaces with rough vertical protrusions of several micrometers.

The apical membranes of some epithelial cells have specialized structures such as microvilli, cilia, endocytic pits, and tight junctions. SICM can provide high resolution images of these structures in live cells. In addition to revealing nanometer-scale morphology, SICM is also a powerful tool to analyze the dynamics of nanostructures of live cells. The dynamics of fine structures in cell membranes is vital for investigating cell function and facilitating the study of important physiological processes. Formerly, these structures could only be observed by scanning electron microscopy or to a lesser extent by atomic force microscopy, but both methods are only suitable for fixed samples which prevent observing the fine structural and functional changes of membranes in live cells. SICM, on the other hand, is suitable for imaging live cell surfaces bathed in electrolytes. Real-time observations of changes in the cell surface can be continuously conducted under normal physiological conditions. SICM has been used to observe directly the assembly of microvillar structures in living renal epithelial cells. The real-time nanoscale topographical images show that multiple microvilli can either form ridges or break into small isolated structures. The height of microvilli can also be rearranged in response to cell volume change (Gorelik et al., 2003, 2004).

With the introduction of hopping mode, large microvilliar structures like stereocilia of hair cells were observed at high resolution (a diameter of 100 nm or less) (Novak et al., 2009). A kinocilium (true cilium) was also visualized in young postnatal auditory hair cells. SICM has also been used to investigate the dynamic changes of tight junctions. In renal epithelial cells, the morphological changes of tight junctions after hypertonic stress was visualized by SICM and used to determine the role of tight junctions in maintaining epithelial monolayer integrity (Zhang et al., 2005). Besides microvilli, cilia, and tight junctions, the endocytic pit is another specialized membrane structure which can be visualized by SICM (Shin and Gillis, 2006; Shevchuk et al., 2008a). Recent studies have shown that this powerful tool can even be used to visualize the process of clathrin-coated pit closure (Shevchuk et al., 2012). In summary, SICM is a very powerful nanotechnology to study specialized membrane structures in live cells. The use of SICM might be extended to the study of the mechanism of cilium formation or loss in the normal and polycystic kidney disease in renal epithelial cells. Recently, my laboratory has established this nanotechnology and has examined microvilli, cilia, endocytic pits, and tight junctions in mouse cortical collecting duct mpkCCD_c14 cells (Figure 2).

Since SICM uses glass micropipette filled with electrolyte, after achieving topographical image by scanning the cell surface the same micropipette can be used for patch-clamp recording of ion channels. In other words, SICM provides the nanoscale navigation for patch-clamp recording of single ion channel activity at defined positions or structures on the cell surface. This so-called high-resolution or “smart” patch-clamp technique was initially developed by Gu et al. (2002). They successfully mapped ion channels in cardiac myocytes: L-type Ca²⁺ channels and Cl⁻ channels are located only in the regions of T-tubule openings. Using the conventional patch-clamp technique, the maxi-anion channel was hardly caught because the “smart” patch-clamp technique showed that the channel is mainly located at the openings of T-tubules and along Z-lines in adult cardiomyocytes (Dutta et al., 2008). After detecting the membrane structures of isolated guinea-pig ventricular myocytes with a lateral resolution of 1.86 μm, a cystic fibrosis conductance regulator-like Cl⁻ channel was precisely localized in these myocytes, which are located both in z-grooves and in the inter-groove region, but not at the mouths of t-tubules (James et al., 2010). As described above, SICM can also visualize specialized membrane structures such as microvilli and tight junctions. The “smart” patch technique makes it possible to easily locate the patch pipette at a region of interest on the cell surface including microvilli and tight junctions to record the ion channel activity (Gorelik et al., 2004; Yang et al., 2010). Since atomic force microscopy showed that the epithelial sodium channel (ENaC) is predominantly located in the microvilli of distal nephron cells (Smith et al., 1997), it would be very interesting to detect how this unique localization is related to ENaC function and regulation.

SICM can be combined with simultaneous confocal fluorescence imaging, resulting in a new technique named scanning surface confocal microscopy. This technique allows investigators to study the relationship between cell surface membrane structures and fluorescently labeled proteins or nanoparticles in both fixed and live samples. SICM can be performed at a sufficient resolution on living cells to image fixed or slowly diffusing individual protein complexes in the plasma membrane and to follow their reorganization over time. Such proteins are likely to be present on highly structured cells where specific functions are associated with particularly specialized regions or domains. The cell membrane exhibits numerous submicrometer-sized surface structures that could be topographically confused with virus particles. This problem can be solved with simultaneous confocal microscopy localization of fluorescently labeled molecules. Using this technique, previous studies showed that virus-like particles were randomly distributed across the cell membrane and had no specific interactions with specialized membrane structures such as microvilli (Shevchuk et al., 2008b). The combination of SICM with confocal microscopy also allows investigators to study nanoparticle uptake. Previous studies showed that nanoparticles with two different sizes (50 nm and 1 μm) can be internalized in equal numbers by human alveolar type-1-like cells (Kemp et al., 2008). In addition to the nanoparticles described above, this technique also can be applied to any virus or other nanoparticle such as gene delivery agents as long as they can be labeled with a fluorescent moiety. It is anticipated that this procedure will be useful in characterizing the earliest interactions of these particles with the cell and contribute to identifying novel drugs to prevent viral infection as well as in the development of gene therapy reagents. This technique should find widespread application for studying the relationship of fluorescently tagged molecules within the cell membrane.

Regulation of cell volume is a fundamental mechanism for cellular homeostasis. Changes in cell shape, movement, and volume reflect normal cellular functions such as secretion, ion transport, and adaptation to environment. In order to study the physiology and pathology associated with the regulation of cell volume and cell membrane movement, an appropriate technique is needed, which allows quantitative and high-resolution characterization without influencing the cell functionality. SICM is one of the most advanced ways to meet these requirements. The technique can measure a wide range of volumes from 10⁻¹⁹ to 10⁻⁹ liter. The cell volume, as well as the cell volume of small cellular structures such as dendrites, processes, or microvilli, can be measured at the 2.5 × 10⁻²⁰ liter resolution (Korchev et al., 2000a; Kemp et al., 2008). Membrane movements in the range of 30 min to a few hours can be quantitatively monitored with SICM at a lateral resolution of 500 nm (Happel et al., 2003; Kemp et al., 2008). Through monitoring the volume of either cells or cellular structures, SICM is a powerful tool to investigate how a functional epithelial monolayer maintains its integrity when given hypertonic stress (Zhang et al., 2005).

Mechanosensitive ion channels convert external mechanical force into electrical and chemical signals in cells, but their physiological function is hard to detect. SICM is a satisfactory physiological tool to stimulate these channels in living cells by means of a nanopipette. This method can be straightforwardly extended to noncontact mapping of both the mechanical properties of the cell and mechanosensitive ion channels (Sanchez et al., 2007). Sanchez et al. produced a non-contact, controlled mechanical stimulation on human and rat dorsal root ganglia sensory neurons and, simultaneously, used whole-cell patch-clamp recordings and measurements of intracellular Ca²⁺ concentration to validate the effects of mechanical stimulation on mechanosensitive ion channels. SICM can provide repetitive mechanical stimulation of a specific cellular region without introducing any damage, and serves as a stable model to study cell mechanotransduction and test novel drugs that may inhibit or modulate mechanosensitive ion channels. When the scanning is performed near the cell membrane, SICM can be used to interact with growing neurons to induce their remodeling (Pellegrino et al., 2009, 2011). Since the guidance of axon growth cones to their targets is essential for establishing neural circuits, the technique can significantly promote neuroscience research.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.