Since the microscope has been invented in the 16th century, microscopes are used to study biological samples with a resolution beyond the limit of the human eye. Today, in general two different types of microscopes exist. The first and larger group comprises microscopes that operate in the far field. These microscopes use light or, in case of scanning or transmission electron microscopes, electrons to illuminate the sample and detect for example reflection, scattering or fluorescence. Since waves are used to illuminate the sample, these microscopes are subject to the diffraction limit that has been described by Ernst Abbe and that links the resolution of the microscope to the wavelength of the radiation used to illuminate the sample [1]. Nevertheless, recent developments in far-field microscopy allow to utilize certain physico-chemical properties of fluorescent molecules to circumvent Abbe's limit [2–6].

The second group of microscopes comprises the scanning probe microscopes. These microscopes use a tiny probe to measure a physical variable that depends on the distance between the probe and the sample surface. The first scanning probe microscope, the scanning tunneling microscope, was introduced in 1982 by Binnig, Rohrer and co-workers. It uses the tunneling current between a conducting probe and a conducting surface to determine positions of equal distance between tip and surface and thus to reconstruct the topography of conducting samples [7]. This technique, which was awarded with the Nobel Prize in 1986, requires the sample to be conductive and furthermore operates in vacuum, and hence can only image biological samples in an artificial environment after complex sample preparation. The second scanning probe technique developed by Binnig, Rohrer and their colleagues, the atomic force microscope (AFM) [8], utilizes the deflection of a soft cantilever with a sharp cone at its end that is dragged over the sample and hence senses its surface structure. Although the strengths of the AFM are the imaging of hard samples at even sub-atomic resolution [9], it had been quickly adapted to image biological samples [10–13]—the state of the art of AFM-imaging of biological samples has been reviewed recently [14]. Despite these successful recordings, the force which is applied by AFM to the sample cannot be neglected [15,16] and hence imaging living cells without biasing still remains a challenging task.

In 1989, Paul Hansma and colleagues invented the scanning ion conductance microscope (SICM) [17]. It monitors the ionic current through the tiny opening of an electrolyte filled glass micro- or nano-pipette. The current flow through the opening is hindered if the tip is in close proximity to a non-conducting surface. This relation between ionic current and tip-sample distance is utilized to determine the topography of the sample, either by calculating the distance between probe and sample from the current flowing through the tip opening, or, more often, by determining positions of equal resistance change which are used to reconstruct the sample topography.

The general assembly of a SICM is similar to electrophysiological setups, as shown in Figure 1A. A voltage is applied between two Ag/AgCl-electrodes, one of which is located in bulk electrolyte solution, the other one is inserted into a glass pipette containing the same electrolyte solution. The leakage current I_L through the opening of the pipette depends on the distance d between probe and sample surface.

The mathematical descriptions of an ionic current through the aperture of a micro-pipette base on the description of the Scanning Electrochemical Microscope by Bard and colleagues [18,19] that was adapted to SICM by the group of Harald Fuchs [20]. The different approaches and approximations to describe the current through a pipette depending on the shape of the pipette have been reviewed in detail recently [21], hence we focus on the description of the imaging process for a given pipette.

Scanning electron micrographs of a scanning probe are shown in Figure 1B. The probe consists of a bulk region (top in Ba) and two tapering regions, the second one magnified in Figure 1Bb. The opening diameter of this pipette approximated one micrometer (Figure 1Bc,Bd). If combined with physiological bath solution, probes with this geometry have an access resistance in the range of 4 MOhm to 6 MOhm. A pipette like this one allows recordings with a resolution in the micrometer range. However, pipettes with a smaller opening diameter allow recordings with resolutions in the nanometer range. Using nano-pipettes with an opening diameter of 13 nm a spatial resolution in the range of 3 nm–6 nm has been observed [22]. However, the resolution of SICM still is a matter of debate, since in contrast theoretical considerations suggest a resolution of 3/2 inner pipette diameters [23].

As shown in Figure 2A, the circuit of a SICM can be described by a series of three resistors. The resistors r_P and r_B denote the resistances of the pipette and the bulk solution, respectively, and depend both on the conductance of the electrolyte as well as on the position and shape of the Ag/AgCl-electrodes (see [21] for review). The resistance r_L of the leakage current I_L through the aperture of the electrode depends, besides the shape of the tip, on the opening diameter and the distance d between the aperture and the sample surface [20,23,24].

The total resistance R of the setup is the sum of these three resistances. Since r_L is a function of the distance d between tip and sample, the total resistance also depends on d: (1) R ( d ) = r P + r L ( d ) + r B .

Note that we neglect that the bath resistance r_B is a function of the position of the tip opening (and therefore of d) since it depends on the distance between tip opening and bath electrode. Furthermore, we neglect the voltage drop at the surfaces of the AgCl-electrodes, assuming they are constant and much smaller than the voltage drop at the pipette tip. Furthermore, we denote the reference resistance of the entire system at infinite tip-sample distance R(d = ∞) = R_∞.

A typical approach curve recorded with a micro-pipette as shown in Figure 1B is shown in Figure 2B. Note that since the initial distance d₀ between tip and sample is unknown, the deflection z of the piezo is recorded instead. The distance is the difference between the initial distance d₀ and the piezo deflection z: d = d₀ − z. While the resistance R(z) of the system matches the reference resistance R_∞ as long as the tip-sample distance is large and hence R(z)/R_∞ ≈ 1, the resistance increases asymptotically as the pipette tip approaches the surface (hence z → d₀ and d → 0).

A normalized approach curve as shown in Figure 2B can be approximated as (2) R ( d ) R ∞ = 1 + D d = 1 + D d 0 − z , where D denotes the tip-sample distance that doubles the resistance [20,25]. The parameters d₀ and D can be determined by numerically fitting this equation to the data. D is determined by the geometric properties of the electrode and the electrolyte. Basing on D, every distance D_ΔT at which the resistance has increased by a certain amount ΔT can be determined by calculating D_ΔT = D/ΔT. Since often recordings of approach curves end before the resistance is doubled, it might be more successful to directly fit (3) R ( d ) R ∞ = 1 + Δ T D Δ T d = 1 + Δ T D Δ T d 0 − z .

Figure 3 shows an approach curve with Equation (3) fitted (red line) to the low-pass filtered data (blue crosses) of an approach curve, ΔT was chosen as 0.03 (indicated as relative threshold T = 1 + ΔT by the black line). For the given pipette, which was similar to the one in Figure 1B, D_0.03 was 437.4 nm ± 16.0 nm (dashed black line). Note that here the approach curve was plotted versus the tip sample distance d, the piezo deflection is indicated by the gray axis at the top of the diagram. The resistance starts to increase approximately at a tip sample distance of 2.5 μm, that is approximately 2.5 inner diameters of the pipette. Nevertheless, a distinct signal that can be clearly distinguished from the remaining noise appears at approximately 1.5 μm (and hence 1.5 pipette diameters).

Since its invention, various ways to determine the sample topography with a SICM have been developed. A brief overview of the imaging modes is provided in Table 1. Although seldom used for imaging, let us start with a general scan mode as depicted in Figure 4A first. Here, the pipette is lowered towards the surface until a resistance change is detected. The pipette is then moved laterally while maintaining its z-position and monitoring the resistance. Assuming a constant conductivity of the electrolyte solution, the distance d between tip and sample can be calculated using Equation (2). However, the distance range in which the probe senses the sample is small (see Figure 3), hence only flat samples with small variances in height can be imaged.

Cells are soft, delicate samples that are easily affected during the imaging process. Even if they are fixed, which means that the proteins within a cell are altered chemically such that their biochemical activity is arrested and the cells are not considered as living, cells remain soft and hence prone to distortions by the scanning probe. Rheinlaender and colleagues compared the influence of the imaging process on fixed cells that were imaged both by AFM and SICM [38] in AC- and tapping-mode. They showed that in AFM images of fine cellular structures such as cell extensions, these structures appear lower and thinner than in the corresponding SICM recordings (diagonal arrows in Figure 7), verified by the comparison of AFM and SICM images of collagen fibrils and chromosomes [39]. Furthermore, by comparing images recorded with opposite scanning directions, they showed that parts of the cell membrane that were only attached loosely to the substrate are shifted towards the scanning direction by AFM and hence appear at different positions (bold arrows and panel h in Figure 7). In contrast, in SICM recordings the locations of those loose structures were independent from the scanning direction (e and k in Figure 7).

Besides these distortions, AFM also irreversibly destroyed parts of the sample, indicated by the horizontal arrow and the dashed circle in the images shown in Figure 7. While these structures were imaged by SICM repeatedly without notable differences (Figure 7c,d), they were not observed by AFM (Figure 7f,g). A SICM scan succeeding the AFM scans did not show these structures either (Figure 7i,j), suggesting that these structures have been destroyed by the AFM imaging process.

Furthermore, the authors compared the vertical approach modes of both scanning probe techniques: force mapping and backstep (hopping/STA) mode. They found that due to the shape of the AFM tip, the basal part of the tip interacts with the sample at steep slopes. This results in artificial topography signals at steep, narrow structures. In contrast, due to the aspect ratio of the probe, SICM allows the recording of these structures (Figure 8a–d). Additionally, the quality of SICM images has been validated by comparing SICM and scanning electron microscopy images [39].

When even higher samples are imaged, the interaction of the basal part of the tip results in the observation of artificial, tail-like structures (dotted region in Figure 8e) that are not observed with SICM (Figure 8f). Furthermore, due to the interaction of AFM tip and sample, the slopes of the sample are smoothed and therefore the sample appears broader than in the corresponding SICM recordings (Figure 8g,h).

In contrast to fixed cells as investigated above, living cells maintain parts of their physiological activity even in culture—as generally used for SICM imaging—and hence, depending on the cell type, migrate, change their shape, contract or show electrical activity. Therefore, imaging living cells is a complex task. The first SICM recordings of living cells have been reported by Korchev and colleagues [40,41]. By applying SICM to A6 cells, Gorelik and colleagues revealed the life cycle of microvilli and how a monolayer of A6 cells maintains its integrity [42–44]. SICM furthermore helped to characterize the molecular identity of the receptors mediating ATP stimulated Na⁺ channel activity in renal epithelial cells [45].

Cardiovascular cells have been extensively studied by SICM [46]. SICM recordings of cardiomyoctes allowed the definition of a new parameter, the z-groove index, to describe the morphology of the cell surface [47]. Furthermore, the differences in failing and healthy ventricular myocytes have been determined [48] and it has been shown, that after prolonged mechanical unloading the surface of cardiomyocytes becomes more flat [49,50].

Since SICM allows detailed, three-dimensional imaging, it has been used to investigate and characterize different cell types or stages of cellular development that can be distinguished morphologically. It revealed that corticosteroids reverse the morphologic effect of cytokines on survival and differentiation on oligodendro-glial progenitors [51]. Furthermore, SICM studies of the neuroblastoma SK-N-SH cell line showed that this cell line contains all three morphologically different cell types known from neuroblastoma [52] and supported the electrophysiological recordings from PC12 cells, which become more neuron-like after exposure to nerve growth factor [53].

SICM has been used to investigate the morphological response of endothelial cells to shear stress [54,55] and to determine the coverage of holes in an artificial membrane that was used as a substrate for astrocytes and fibroblasts [56].

Due to its resolution beyond the limit of conventional light microscopes, SICM has been applied successfully to investigate surface changes after artificially induced exocytosis and revealed that the membrane of a fraction of 16 % of the investigated cells showed tiny dips shortly after exocytosis [57]. Furthermore, SICM has been applied to determine the toxicity of various nanoparticles that induce holes in the cell membrane [58,59]. On an even smaller scale, SICM has been used to determine the movement of protein complexes within the cell membrane of living spermatozoa [22].

Although the investigations detailed above have only been possible because of the nearly non-invasive imaging characteristics of SICM, the scanning probe can be easily used to stimulate the sample. The probe can be used to apply a force on the membrane, allowing the electrophysiological recording of mechano-sensitive currents [60] or pressure applied through the scanning pipette can be used to determine the mechanical properties of the cell membrane [61,62]. Both, force by the interaction of pipette and membrane and pressure through the pipette can be used to direct growth cones of leech neurons [63,64]. Note that even the flow of electrolyte through the pipette opening that is not compensated by the capillary tension might stimulate live cells and thus impair the sample, particularly if relatively large pipettes are used.

Since probe microscopy images reveal the three-dimensional shape of a cell, not only the cellular surface but the cellular volume can be investigated with SICM [26]. Furthermore, volume changes can be attributed to specific regions of a cell [36] or, after specific processing of the data, be divided in frontal and rear volume changes even in cells that change shape and position between succeeding scans [65], as shown in Figure 9. If the resolution is decreased and a scanning time of approximately two minutes is achieved, fast volume regulatory processes can be investigated with SICM [66]. An exemplary recording is depicted in Figure 10, showing the typical fast regulatory volume increase (RVI, red arrow) after application of hyperosmolar solution and regulatory volume decrease (RVD, orange arrow) after switching back to lower osmolar control solution.

SICM measurements comprise two separated volumes, the bulk solution and the solution within the scanning probe, both connected only by a small interface. Since SICM allows both the determination of the sample topography and the precise control of the tip-sample distance, it has been widely used to deposit certain molecules onto a surface [82–87]. By trapping [88] fluorescent dye molecules in the tip of the probe the probe was applied as a fluorescent nano-sensor [89]. Furthermore, the scanning probe has been used to spatially modify the pH or Na⁺ concentration by the delivery of corresponding ions [90] and has been used to pipet femtoliter-sized droplets [91]. A further application of SICM is the investigation of nano-pores within an artificial membrane [92–95]. Two reviews detail these more chemical applications of SICM [96,97].

Scanning electrochemical microscopy (SECM) is a microscopy technique used to spatially resolve analytes by determining faradayic currents from redox reactions. The close relationship of SICM and SECM has been described in a review of the use of SECM in neuroscience. For more information about SECM the reader is referred to this review and the references therein [98].

To combine SICM and SECM measurements, the scanning probe has to be modified to determine faradayic redox currents. Various probe preparation techniques allowed the simultaneous recording of the topography and an analyte at increasing resolutions [99–103]. It can even be combined with the delivery of molecules through the pipette [104] and used to investigate the transport of redox species through pores in an artificial membrane [105]. Recently, it was applied to detect neurotransmitter release from living hippocampal neurons [106].

Here we reviewed the various modes of SICM for studying biological samples. As shown for various examples, the strength of the method particularly resided in its electrical distance control avoiding mechanical damage of the delicate membranes of living cells. We detailed the combinations of SICM with other micro- or nanoscale detection methods such as fluorescence microscopy, patch clamp and ion selective micro-electrodes.

We conclude that although further refinements particularly regarding the temporal resolution are required, that SICM is an emerging technique for life scientists in addition to further potential applications ranging from material sciences to medicine.