INVITED REVIEW
Pushing, pulling, dragging, and vibrating renal epithelia by using atomic force microscopy

Robert M. Henderson1 and Hans Oberleithner2

1 Department of Pharmacology, University of Cambridge, Cambridge CB2 1QJ, United Kingdom; and 2 Institut für Physiologie, Westfälische Wilhelms-Universität Münster, D-48149 Münster, Germany


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

Renal physiologists focus on events that take place on and around the surfaces of cells. Various techniques have been developed that follow transport functions at the molecular level, but until recently none of these techniques has been capable of making the behavior of molecular structures visible under physiological conditions. This apparent gap may be filled in the future by the application of atomic force microscopy. This technique produces an image not by optical means, but by "feeling" its way across a surface. Atomic force microscopy can, however, be modified in a number of ways, which means that besides producing a high-resolution image, it is possible to obtain several types of data on the interactions between the ultrastructural components of cell membranes (such as proteins) and other biologically active molecules (such as ATP). In this review we describe the recent use of the atomic force microscope in renal physiology, ranging from experiments in intact cells to those in isolated renal transport protein molecules, include examples of these extended applications of the technique, and point to uses that the microscope has recently found in other areas of biology that should prove fruitful in renal physiology in the near future.

kidney; scanning probe microspcopy; Madin-Darby canine kidney cells; ion channels; nuclear membrane; plasma membrane


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

THE ATOMIC FORCE MICROSCOPE (AFM) was developed in 1986 (7). Like the scanning-tunneling microscope (STM) that preceded it, the AFM is a scanning probe microscope. These instruments differ from "conventional" optical and electron microscopes because they work by moving a probe back and forth across a surface and recording features as the probe encounters them. Scanning probe microscopes thus produce images that are not compromised by the limitations of the wavelengths of the various types of electromagnetic radiation. This means that very high resolution can be obtained. In some cases, studying physically hard samples, down to atomic resolution, but with softer samples the maximum resolution is at present on the order of 1 and 0.1 nm in the lateral and vertical directions respectively. The probe of the STM is made from a conductive material, and the instrument works by measuring a current, the "tunneling current," between the probe and the sample. Thus the STM is suitable for use only to study electrically conductive materials. The AFM, on the other hand, can be used on nonconducting specimens. As a result of the high resolution and the fact that it can be used on samples under fluid, the AFM (which was originally designed with applications in physical sciences in mind) soon attracted interest among biological scientists (26) as a tool for imaging cellular and subcellular structures under physiological or near-physiological conditions and to study dynamic features of molecular behavior at high resolution in "real time" under these conditions. Biological work using the AFM has shown considerable growth throughout the 1990s (23). As a result partly due to technical innovations, the realization that useful images could be obtained under fluid, together with the ready availability of microscopes from a number of manufacturers, the field has moved on and expanded far beyond that envisaged when the instrument was developed. Not surprisingly, the potential of the AFM was swift to attract interest among renal physiologists, and the use of the technique in renal cellular physiology has, to a great extent, parallelled the use of the AFM in cell biology in general. In this review we will outline the use that the AFM has found in renal physiology and, because AFM technology is continuously evolving, describe mechanisms in renal cells that have been revealed by the AFM and outline recent developments that show promise for future studies on renal cellular function. Coverage of the whole field of biological ATM is outside the scope of this review, but readers who wish to explore further topics are directed to a number of recent more comprehensive works (8, 11, 16, 18, 22, 34, 35, 40, 42, 50, 51, 69, 78, 81).


    OBTAINING IMAGES AND FORCE DATA WITH THE AFM
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

The AFM probe is typically made from a pyramidal crystal of silicon nitride (Si3N4) deposited onto a gold-coated, flexible cantilever. A sample is prepared on a flat substrate and moved so that it makes contact with the probe. The sample is then moved back and forth in a raster pattern, and the probe is deflected vertically as features in the sample move under it (Fig. 1). The movement is controlled by a series of piezoelectric drivers, and the control is such that the probe can be positioned in either horizontal x- or y-dimensions or in the vertical z- dimension very accurately (in the subnanometer scale). In some microscopes, especially those designed for use with biological specimens, (e.g., Ref. 25), the probe is mounted on the bottom of the piezo assembly and is itself moved across the sample, which remains static. This means that the sample can be simultaneously observed by using an inverted optical microscope and the AFM. With both configurations of the AFM, a low-powered laser is focused onto the cantilever and is reflected onto a series of photomultiplier detector elements. As a result, when the probe scans across a surface and meets some sort of obstacle, the cantilever is deflected, changing the reflected angle of the laser, and therefore affecting the signal detected by the photomultipliers. The photomultipliers' signals are fed into a computer, which then constructs a three-dimensional image from the information received. The principle of the AFM is therefore quite simple, and its implementation is dependent on the availability of suitable sensitive photomultipliers and piezo control mechanisms.


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Fig. 1.   Fundamental elements of operation of atomic force microscope (AFM). A low-powered laser is focused onto top of cantilever (which bears the probe). The sample, on a suitable substrate, is then scanned by probe in a raster pattern, and tip deflection is recorded by altered reflective angle of laser that is detected by a photomultiplier array. The signal from the photomultiplier array is processed by a microcomputer to produce an image of sample, and the computer also provides a feedback signal to AFM piezo drivers to control the force of interaction between the probe and substrate. In most microscopes the probe remains stationary whereas sample is scanned back and forth on a piezo-driven stage, but in some microscopes the reverse is true. The probe is attached to piezo drivers, and substrate remains stationary. For more details, see text.

In constructing an image with the AFM, the probe normally has to make contact with the specimen. This means that as the probe moves across it can distort the sample (and thus produce misleading images) because it normally applies both vertical and lateral force. This is particularly important when using AFM on biological specimens (which are relatively soft compared with many specimens studied in the physical sciences), and it is therefore important that the force applied to the sample by the probe can be minimized. This is partly achieved by the cantilevers, on which the probes are mounted, having very low spring constants (generally in the region of 1 N · m-1, similar to the intermolecular forces producing attraction between biological macromolecules). This means that as the probe moves across a sample it can be deflected vertically by features it might encounter without the danger of the cantilever being so stiff that it sweeps away all obstacles in its path. In addition, the force can be reduced by fine adjustment of the microscope at the start of an experiment. This is achieved by means of a "force curve" (Fig. 2). To construct the force curve, the probe is held stationary (in the horizontal directions) on the substrate, and the tip is oscillated vertically. As the probe and substrate make contact (during the downward-moving "approaching" phase of the probe), the cantilever bearing the probe is deflected, and this deflection is registered by the photomultipliers. As the probe and substrate are drawn apart (during the "withdrawal" phase) the cantilever is again deflected, returning to its original position, but often being further deflected as a consequence of the probe "sticking" to the substrate, a result of the adhesion forces (which may be chemical, electrostatic, or even magnetic) between probe and substrate. These adhesion forces are a consequence of the way in which the AFM works. By examination and by adjustment of the force curve, they can be minimized in such a way that excessive vertical forces are not applied to the sample. In addition, adhesive forces between probe and substrate are reduced considerably if imaging is conducted under fluid, a medium that is often to be preferred in biological imaging. Besides adjustment of the microscope to optimize image generation, the force curve also provides a useful measure of the degree of attraction between probe and substrate and may thus be used to distinguish different areas of the same sample if they have differing physical characteristics. This feature is exploited by using the technique known as "force volume" imaging. Here, force curves are generated at a number of points along each raster line, and an image is built up that effectively produces a map of probe-substrate interactions across the surface of a sample at regular intervals, which is displayed together with the conventional AFM image.


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Fig. 2.   Examples of force curves contrasting situation with a sample in air (top) with that under fluid (bottom). Essential details are given in text. Probe is held stationary over substrate and then oscillated up and down ("extended" and "retracted"). At A, probe is not in contact with any substrate and so no deflection is registered. At B, probe meets substrate, and at C it is advanced farther downward onto substrate and so cantilever bearing probe is deflected. This is shown in "y"-axis of curve. Piezo drivers then begin to withdraw probe upward (retract). Because probe and substrate are physically attracted, they maintain contact (D), even when probe has been withdrawn before point where it originally made contact with substrate. At E, probe loses contact with substrate and jumps back to its original position (F). The important point concerning the force curve is that degree of attraction between probe and substrate must be minimized. The measure of this attraction is given broadly by size and shape of triangle lying below dotted line in 2 diagrams. Forces are minimized when recordings are made under fluid, which is the preferred environment for biological experiments.

The AFM may be used to obtain data by using various different "modes" of recording. These are described below and outlined in Table 1. The "conventional" use of the AFM described above is known as "contact mode." The probe applies constant force while it scans, and the tip and piezoelectric drivers are connected via a feedback loop circuit to move the probe and substrate toward and away from each other to maintain the constant force. This leads to a representation of the height of topographical features in the image obtained ("height mode" recording). Because the feedback loop is driven by changes in cantilever deflection, it is also possible to monitor these deflection signals and use them to construct an image. This type of data collection is called "deflection mode" recording, and it typically produces images in which edges of features are emphasized (which can be useful under some conditions) but gives no quantitative data on height. Contact mode recording, with either or both height and deflection images, is very effective under many circumstances, but even when vertical forces are minimized by using the force curve, the application of lateral force as the probe is drawn across the surface of the sample is unavoidable, and this can also distort the image produced. If the force applied is excessively high, the sample can even be damaged or perhaps pushed around the substrate by the probe, which behaves like a small snowplough. If the applied force is too low, then the probe can bounce off the sample as it moves over topographical features. These limitations can be overcome by setting the tip oscillating vertically at a high frequency while scanning normally in the x-y direction, a mode of recording known as "tapping mode" (Fig. 3). The reflected signal from the cantilever detected by the photomultipliers then describes a sine wave, the amplitude of which is altered as the probe moves over any physical feature on the surface. The tapping mode technique results in much lower lateral (and also vertical) forces being applied to the sample compared with those seen in contact mode recording and produces concomitantly clearer images with less scope for distortion of samples. Tapping mode was originally only possible on dry samples imaged in air or a vacuum, but even under these conditions the adhesive forces between a dry sample and the probe can be such that distorted images can result. To circumvent this problem the technology has been developed that makes it possible to make tapping mode recordings under fluid (24, 66).

                              
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Table 1.   Comparison of different modes of data acquisition with the AFM



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Fig. 3.   Illustrating the principle of "tapping mode" (A and B) and "phase imaging" (C and D). In tapping mode the probe is set oscillating, describing a sine wave (A). Amplitude of oscillation is altered when probe meets any perturbation on substrate (B), and change in amplitude is used to produce an image. In phase imaging, tapping mode is used, but behavior of probe in comparison to that predicated by the signal driving its movement is monitored. In C, probe moves according to signal produce by driver (sine wave). However, in D, tip is physically attracted to gray area of substrate, and thus there is a lag between the expected oscillation and that observed. This phase difference can be recorded and give a measure of degree of attraction between probe and substrate.

A further development of tapping mode recording has recently become available. This is called "phase imaging" (5, 46). In tapping mode recording, the cantilever is oscillated at high frequencies (~300 and 12 kHz in air and fluid, respectively). If the cantilever is oscillating freely in air then the waveform signal detected by the photomultiplier should be exactly in phase with the signal driving the piezoelectric element producing the oscillation. However, if the probe comes into contact with some substrate, any attractive force between the probe and substrate will tend to cause the tip to be held back in contact with the substrate, and so the photomultiplier signal will become out of phase with the piezoelectric driver's waveform. This phase lag can be used to construct an image for the AFM that gives a measure of adhesive forces between the probe and substrate, and this mode of recording constitutes phase imaging. The technique is promising because it provides information on probe-substrate interactions that need not necessarily be reflected in apparent changes in topographical features of the sample.

Although there are a number of different ways in which images can be obtained in AFM, they are all dependent on the types of probes used. A consideration therefore arises in choosing the type of tips on probes used in imaging experiments. Although for most purposes a tip with a conventional pyramidal profile produces good images and force information, various types of other probes with tips of different aspect ratios can be produced. Under certain circumstances it is important to use probes that have a sharper profile, effectively tapering at a sufficiently acute angle at the tip so that they are able to penetrate small crevices in the sample. If unable to do so, fine details of the structure may be lost in the image. Various techniques have been adopted to optimize tip profiles. Probes are commercially available that have been sharpened or that have sharply tapering columns of Si3N4 deposited at their ends to improve resolution, but the sharper the tip, the more difficult it is to make, and this is reflected in the price. A recent development of potential importance has been the production of Si3N4 probes that have had carbon "nanotubes" attached to their tips (12). These nanotubes are long (up to 1 µm) and thin (1-5 nm in diameter at the tip), optimizing image production by their ability to gain access to small recesses in surfaces without exerting large forces. In the future it is hoped to refine the nanotube technique to produce tubes tipped with a single hemifullerene dome [essentially a bisected molecule of buckminsterfullerene (12)].


    THE AFM AND INTACT RENAL CELLS
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

The use of the AFM in renal physiology has been the subject of a number of reviews (e.g., 29, 46, 54, 56, 59, 61). In the early years of the AFM, experiments tended to concentrate on imaging intact renal cells. These experiments continue but are now complemented by studies on subcellular structures. Part of the reason that renal cells have been the subject of interest to workers using the AFM has probably lain in the ready availability of stable renal cell lines and the interest in membrane transport functions possessed by renal cells; the AFM is an ideal instrument for examination of membrane structures. As a result a substantial body of work by a number of different groups has been produced by using Madin-Darby canine kidney (MDCK) cells, CV-1 African green monkey kidney cells, A6 cells from toad kidney, and also opossum kidney (OK) cells. An image of the surface of a living OK cell is shown in Fig. 4. These studies have provided valuable data but also perhaps served to highlight the difficulties of using the AFM in intact cells. Hoh and Schoenenberger (33) reported that, by using relatively high scanning forces of >2 nN, numerous features could be identified in living cells. High scanning forces (in the nN range) have proved to be a requirement when intact cells are being imaged (28). Although (33) many structures were evidently located on the cells' surface, others such as the nucleus and cytoskeletal elements were clearly inside the cell. The identification of these submembrane features was dependent on the probe of the AFM applying force to deform the cell membrane, and thus "feel" the structures beneath. This suggestion was confirmed by the authors' application of glutaraldehyde to the cells while the probe remained in contact with the cells. The fixative led to a stiffening of the cell and reduced capacity for deformation of the plasma membrane. In a closely parallel study (43) Le Grimellec et al. produced better resolution in images of fixed cells, and an improvement in resolution of the images of living cells was also obtained by enzymatic treatment of the cells' surfaces to remove the glycocalyx. Under these circumstances, the authors were able to distinguish numerous protrusions from the cell membrane that they identified as proteins, through treatment with proteases, although any further characterization of these structures proved difficult. This is partly due to the image being compromised by deformation of the protein structures produced by the large forces applied by the probe and also to the presence of large numbers of microvilli on the surface of the cells that, in turn, are deformed by the probe (46). The situation has been studied in some detail in glial cells (72) by the high forces applied to cells, but the images were further compromised by cellular components sticking to the probe as it passed over the cells. These problems highlight a major concern, because the lack of ability to routinely image and identify subcellular structures compromises the usefulness of the AFM in cell biology. The situation might be expected to be partly resolved by the development of tapping mode recording that can be used on samples in fluid. Taking advantage of this, Le Grimellec and co-workers (44) used tapping and contact mode recording. Subsequently, Le Grimellec et al. (45) showed that by using small scan sizes and carefully adjusting the instrument it is possible to gather images of membrane structures in the same cell by using low forces (down to 20 pN), although the technique would appear to be exacting.


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Fig. 4.   AFM imaging of living, intact renal epithelial cells. Shown is surface topography of cells in an 11-day-old culture of opposum kidney cells grown on a glass coverslip. Cells were imaged in Hanks' medium by using tapping mode AFM. Cell surface appears very irregular, with long microvilli either isolated (black arrow) or organized in bundles (white arrow). Horizontal and vertical scales are indicated. Image is kindly provided by Dr. C. Le Grimellec and Dr. V. Vié of L'Institut universitaire de Recherche Clinique, Montpellier, France.

Further interest in intact kidney cells has been focused on imaging dynamic properties of MDCK cells (57, 77). In one of these studies (77), dynamic movements of the cell membrane and associated structures were identified in normal and oncogene-transformed MDCK cells by using time-lapse images obtained with the AFM in contact mode. Some of the movements in the transformed cells were seemingly initiated by disturbance of the cells by the AFM probe. In another study using MDCK cells transformed by alkaline stress [MDCK-F cells (57)] the authors showed that cells were able to move across a substrate by a process of invagination of cell surface, producing endocytosis near the leading edge, which was strongly indicative of the internalization of plasma membrane. The "dimples" produced as a result of the invagination showed a diameter of ~90 nm. In this study quantification of the number and rate of production of endocytic invaginations corresponded closely with the amount of membrane that was necessary to transfer back into the cells to produce cellular locomotion.

The difficulty of identification of specific proteins among a large heterogeneous population on a cell surface can be addressed by using antibodies conjugated to colloidal gold. This technique has been used to study A6 toad kidney cells. In this study the authors labeled the cells with antibodies generated against an amiloride-sensitive epithelial Na channel (ENaC) purified from bovine renal medulla, that had been conjugated to 8-nm-diameter colloidal gold particles (79). The authors were able to identify areas on the apical microvilli of the cells that showed a marked increase in height compared with control. This they interpreted as being due to binding of the antibody-gold particle complex, and they noted that this suggested that the ENaCs are confined to the apical microvilli in this cell type. In fact, the use of colloidal gold (or similar) particles conjugated to antibodies has been used to label and identify various structures for study with the AFM. The molecules investigated include CD3 on human lymphocytes (53, 65); the nuclear pore protein gp62 and nuclear lamin LIII in Xenopus laevis oocytes (73); and Thy-1 antigen distribution in isolated mouse thymocytes (80). The technique is clearly useful, but information about molecular structure is limited because the necessity of attaching an antibody to identify proteins, together with a relatively large gold particle conjugated to a secondary antibody, leads to a masking of the proteins' features. However, such problems could be circumvented by performing paired experiments: first scanning the membrane with putative proteins in the absence and then in the presence of antibodies. Comparison of the two images would reveal not only the spatial distribution of the proteins on the cell membrane, but also some structural information concerning the native "antigen" protein.

Besides investigations of morphology and dynamic events in intact living cells by using AFM, the technique can also be used to gain useful insights into other aspects of living cells' functions, including measurements of cellular elasticity [for example in MDCK cells (1) and platelets (68), and cell volume (e.g., 76).]


    IMAGING SUBCELLULAR MEMBRANE STRUCTURES
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

Although the difficulty identifying specific structures on the surface of an intact cell (where many similar looking structures coexist) may be circumvented by the use of antibodies in paired experiments, the quality of images obtained is also often compromised by the physical flexibility of the cell. Even with the low scanning forces used in tapping mode recording, structures can be pushed below the cell surface by the scanning probe. To address this problem, sections of membrane can be isolated and supported on a rigid surface, thus allowing imaging of the membrane without distortion. This approach has been taken by Lärmer et al. who, studying MDCK cells, were interested in the structures present in the membranes that are removed with the pipettes in the so-called "cell-free" or "excised" mode of electrophysiological patch-clamp recording (41) (see Fig. 5). The authors transferred excised inside-out patches onto poly-L-lysine-coated mica and studied them both dry and under physiological conditions. The authors found that, in contrast to their earlier experiments on intact cells, where it was difficult to detect plasma membrane proteins, resolution in the excised patches was increased considerably with lateral and vertical resolutions of 5 and 0.1 nm, respectively, making it possible to see proteinaceous protrusions from the membrane (the structures could be degraded by the action of pronase). The improved resolution made it possible to measure the protein structures and to estimate the molecular weights of the proteins. These were between ~50 and 710 kDa (with a median value of 125 kDa). The proteins had a distribution in the plasma membrane patch of ~90 particles/µm2 of membrane. The same group (13) has used a similar approach to examine nuclear pore complexes. These have the advantage of being large structures that have previously been studied extensively by using electron microscopy (for review, see Ref. 62). It was shown that complete nuclear envelope together with nuclear pore complexes (NPCs) could be excised from the nucleus after their functional identification in patch-clamp experiments (13). The fact that NPCs are easily identifiable has meant that they are ideal membrane structures for the application of the AFM, and they have been the subject of several studies by a number of groups (e.g., 10, 15, 19, 20, 55, 63, 70). An image is shown in Fig. 6.


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Fig. 5.   A: patch from Madin-Darby canine kidney (MDCK) cell imaged under fluid. Image clearly shows vesicles under apical surface of membrane, confirmed by profiles in B. In C a higher magnification is shown. Three vesicles were chosen, and profiles are shown in D. They have a height (equivalent of diameter) of ~200 nm. This is consistent with their being subapical vesicles. This indicates that by using patch clamp the membrane can be removed together with cytoplasmic material associated with the inner face of the plasma membrane. This opens an interesting perspective: a patch seems to be more than a lipid bilayer and may often maintain its own microenvironment. This could go some way to explain possibly controversial results in patch clamping, including rundown phenomena, unstirred layer problems, and so on. Images obtained by J. Laermer.



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Fig. 6.   Cytosolic surface of nuclear pore complex (NPC) of nuclear envelope of MDCK cell. Left: 8 subunits (1-8). In the center is the so-called "transporter" (arrow). Right: higher resolution image of transporter. This is a putative protein structure that is thought to belong to NPC machinery. This transporter is highly dynamic. It can open and close and is in charge of macromolecule transport (19). Structure seems to be regulated by aldosterone because, in the absence of the hormone, proteins accumulate on NPC surface (19), and, in the presence of the hormone, proteins are gone (and it is assumed they are being transported through this structure).

Experiments have shown that it is possible to manipulate the nuclear pore complexes by using various physiological stimuli. In experiments on isolated nuclear membranes of Xenopus laevis oocytes imaged under fluid in near-physiological conditions, the AFM has shown that application of ATP leads to a change in the conformation of the NPCs (70). Another group showed opening of the NPC pore to be dependent on the state of intracellular Ca2+ stores (63). In a series of papers describing experiments on the nuclei of MDCK cells, aldosterone treatment for 6 h increased the number of NPCs by 32%. In the same study electrophysiological measurements showed a rise in the nuclear electrical conductance by 39%, concurrent with the increase in NPC number, whereas the nuclear membrane potential also changed from approximately -3 to -6 mV (55). A more detailed examination of the influence of aldosterone on the structure of NPCs of MDCK cells (19) showed the hormone to produce a change in the NPCs from an inactive to an active state (indicated by the active pores showing clear openings between their subunits, in contrast to the inactive and "covered" state). Experiments with the AFM show that TATA binding protein (TBP) molecules (which are required for RNA polymerase II-mediated transcription) attach to and accumulate on the NPC's cytosolic side (10). Electrophysiological experiments using isolated cell nuclei of cultured MDCK cells revealed that TBP translocates into the cell nucleus in the presence of ATP, transiently plugging the NPC. TBP forms multimers in solution (36). Using the AFM (58) to study the protein has directly shown that multimers of TBP are dissociated into oligomers in the presence of ATP and under these conditions they can be translocated into the nucleus.

Thus the experiments on NPCs have shown that ATP can influence the behavior of subcellular structures. The advantage of NPCs in these AFM studies is that they are relatively large, and their structure has been well characterized by using electron microscopy. This means that they provided a good model on which to test the capabilities of the AFM to investigate physiological effectors of protein function. The experiments in (41) have shown that it is also possible to study intracellular surfaces of patches of plasma membrane (which contain numerous protein structures) with the AFM. In these experiments, ATP was shown to produce a change in the NPC [which is a relatively large complex with a molecular mass of ~124 MDa (71)].

This finding opened the possibility that it might be feasible to use the AFM to study the effect of ATP on the much smaller protein structures in the plasma membrane (which have an average molecular mass of 100 kDa). This approach was taken by Ehrenhöfer et al. (14). These authors "glued" apical plasma membranes of MDCK cells to mica with the K+ channel blocker iberiotoxin, a positively charged toxin molecule that binds with high affinity to both plasma membrane potassium channels and mica. The technique involves growing MDCK cells on a conventional substrate (where they align with their basolateral membranes apposed to the substrate). A mica sheet coated with iberiotoxin is then pressed down onto the cells whereon the toxin binds with the apical K+ channels. As the mica sheet is lifted it removes the apical membrane alone, and the cytoplasmic face is then accessible to examination with the AFM. The study (conducted under a fluid, the composition of which mimicked a normal intracellular ionic environment) showed numerous protrusions from the membrane (from 1 to 20 nm in length), arranged in clusters, that were identified as proteins by the effects of pronase and could be stimulated to change shape by addition of ATP (Fig. 7). The authors concluded that plasma membranes are arranged containing "functional clusters" of proteins in the native environment [as earlier postulated by Al-Awqati (2)] and that the "physiological" and physical arrangement of the protein molecules within a cluster required ATP.


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Fig. 7.   Images of cytoplasmic face of plasma membrane of MDCK cell showing effects of ATP. For details see text. The study (conducted under a fluid the composition of which mimicked a normal intracellular ionic environment) showed numerous protrusions from membrane (from 1 to 20 nm in length; arranged in clusters) that were identified as proteins and could be stimulated to change shape by addition of ATP. This is demonstrated by changes in shapes of structures seen in 10 min subsequent to addition of ATP. From Ehrenhöfer U et al. Cell Biol Int 21: 737-746, 1997. Copyright (1997) Academic Press, Ltd.


    ARTIFICIAL LIPID MEMBRANES AND RECONSTITUTION OF MEMBRANE PROTEINS
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

Despite the advances described above, it is still difficult to identify specific membrane proteins in the native environment. The way ahead in addressing the problem of examining structure and behavior of membrane proteins probably lies in producing purified proteins in sufficient quantity and then reconstituting them into suitable environments, ideally reflecting the native plasma membrane. On the other hand, where abundant quantities of proteins are available, they can be prepared in their native environment for imaging with excellent results. Most of this work has been conducted on membrane proteins from bacteria (which can be obtained in large amounts and form two-dimensional crystals). Several papers review this topic (51, 52). The goal for scientists working on eukaryotic membrane proteins is to replicate the conditions provided by the prokaryotes' membranes. In search of this goal, considerable interest has been shown in imaging and manipulation of supported artificial lipid bilayers by using the AFM. In a sense, this has grown out of a large body of work on Langmuir-Blodgett films (for example 9, 37, 85).

Artificial bilayers can be produced by spreading lipids onto a suitable substrate (like mica) (6), and these can be manipulated, defects being "smoothed out," by the AFM. In fact, the degradation of such supported membranes by phospholipase A2 has recently been demonstrated (21). Progress in reconstitution of purified proteins into such artificial membranes remains frustratingly slow because it is notoriously hard to express suitable membrane proteins in sufficient quantities. The recent purification and crystallization of the KcsA K+ channel from Streptomyces lividans (which has some similarity to eukaryotic channels) is a promising development (27, 47).

The closest the AFM has come to producing high-resolution images of renal membrane proteins in their native form is with the water channel aquaporin-1 (AQP1). AQP1 is expressed in the nephron and in high concentrations in erythrocyte cell membranes from which it can be purified in sufficiently large quantities to produce two-dimensional crystals in the presence of phospholipids. Walz et al. (82) have produced processed AFM and electron microscope images of freeze-dried, metal-shadowed crystals of AQP1 that show a tetrameric arrangement, with external protrusions from each subunit, the tetramer having an internal central pore (see Fig. 8).


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Fig. 8.   Image of native aquaporin-1 (AQP1) 2-dimensional crystal. a: overview was recorded in buffer solution at loading force of ~0.2 nN, scan frequency 4.7 lines/s (512 pixel). AQP1 crystal is fragmented, exhibiting cracks and holes where thickness of layer is determined (e.g., in circle). b: at high magnification height signal displays major tetrameric protrusions of 1- nm height. c: in diffraction pattern calculated from the area shown in a, diffraction spots up to reciprocal lattice order 2.9 can be discerned, corresponding to a resolution of 1.04 nm. d: correlation average reveals mainly 1-tetramer/per unit cell, whereas the other tetramer is represented by a pronounced depression with windmill-shaped peripheral protrusions of 0.5-nm height. Bars: 100 nm (a); 20 nm (b); 2.5 nm (c). Side length of d corresponds to 19.2-nm. Image kindly provided by A. Engel (Biozentrum, Basel, Switzerland).


    ISOLATED MEMBRANE PROTEINS
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REFERENCES

In the absence of abundant quantities of purified proteins from renal membranes it is necessary to adopt rather indirect approaches to use the AFM to study the small amounts of available protein. Using a fusion protein of glutathione S-transferase (GST) with the renal K+ channel ROMK1 (32) [which had been prepared to raise an antibody for Immunolocalization of the channel in the kidney (48)], Henderson et al. (30) identified dimeric particles of appropriate size (75). The dimer presumably reflected the normal conformation of GST (49). The experiments were performed with the protein immobilized on mica, in an either dry or aqueous environment (Fig. 9). On cleaving the GST from the ROMK1 with thrombin, single molecules in aqueous solution remained attached to the mica substrate but appeared to aggregate. Addition of ATP to the solution produced a reversible change in height of the aggregates, suggesting that ATP induces a structural change in the ROMK1 protein. ROMK1 is thought to bind ATP and is susceptible to phosphorylation by the action of protein kinase A (PKA) (32). In subsequent experiments, ROMK1 was attached to the tip of an AFM probe and held immobile above a mica sheet in an aqueous solution (60). The channel protein was therefore sandwiched between the tip and mica. Any structural alterations of the sandwiched molecule were transmitted to the cantilever, and this was represented as a change in height registered by the AFM. When the bathing solution contained the catalytic subunit of PKA and 0.1 mM ATP, stochastically occurring height fluctuations in the ROMK1 molecule were observed. The movements were pH sensitive and could be modulated by addition of an antibody to ROMK1 (Fig. 10). The temporal resolution of the structural activity of the ROMK1 protein was shown to be ~1 ms, which means that separate measurements of the protein height were recorded at a high frequency, once every millisecond. An earlier study used a similar approach to investigate conformational changes in lysozyme molecules using the AFM (67) . The measurement of structural changes of membrane proteins in the millisecond range may allow a direct correlation between the protein's function analyzed by electrophysiological techniques and the structural changes in the same protein analyzed by using the AFM.


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Fig. 9.   Shown are images (and cross sections) of ROMK1-glutathione S-transferase fusion proteins. a: Recorded dry, in contact mode. b: Recorded in tapping mode under fluid. From Henderson RM et al. Proc Natl Acad Sci USA 93: 8756-8760, 1996. Copyright (1996) Natl Acad Sci USA.



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Fig. 10.   Demonstration of resolution produced by "molecular sandwich" technique (60). a: Deflection of a noncoated tip with high temporal resolution. b: tip coated with ROMK1. c: dependency on pH of vertical movements produced by ROMK1 protein. Experiments were performed in presence of 0.1 mM ATP and catalytic subunit of protein kinase A (20 mU/ml). See text for details.


    SPECIAL TREATMENT OF TIPS
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ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

The coating of tips of AFM probes with ROMK1 leads to the subject of the coating of tips with other substances, and the usefulness of this technique in renal physiology. There has been a considerable body of work with the AFM concerning the interaction of streptavidin and biotin (4, 17, 83), with one-half of the streptavidin-biotin ligand-receptor pair being used to coat an AFM tip, and the other half of the pair being attached to the substrate. These studies have pointed to the potential of the AFM for measurements of interactions between a large number of biologically important molecules. The techniques used to study steptavidin-biotin interactions have been modified to investigate antibody-antigen interactions and to allow localization of sites on biological surfaces (31, 84). The application of tip-coating techniques is, however, complicated. It is necessary for the ligands to attract each other strongly enough to allow a change in force to be registered during the scan, but they must not attract each other so strongly that the tip is interrupted in its progress over the substrate or that undue distortion of the sample takes place. Furthermore, the binding and unbinding of ligand-receptor pairs unless covalent will be a dynamic process. In the case of a ferritin/anti-ferritin antibody pair, the ligand-receptor binding is in itself a dynamic process that is dependent on the environmental conditions (3). Such methodologies are nevertheless of very great importance, because the demonstrated ability of the AFM to measure interaction forces between ligand-receptor pairs means that it is possible to use force volume imaging to construct a force map across a substrate. This maps the interaction between a molecule adsorbed to the tip and another one on the substrate (15). The forces change as the ligand-bearing tip passes over samples of its complementary molecular pair on the substrate. This approach may be applied to proteins in their native plasma membranes by using coated tips. It has recently been used to identify "hot spots" of localized ATP release from the plasma membrane surface of epithelial cells. In this study the tips were coated with ATPase S1 (which is a subfragment of myosin). As the tip passed over the cell surface, the ATPase hydrolysed ATP on the cell membrane, which produced a disturbance in the normal scanning process of the cantilever. This appeared as a feature in the image produced, presumably registering a conformational change in the ATPase coating the tip (74).


    THE SCANNING ION CONDUCTANCE MICROSCOPE
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ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

A recent development that holds out great promise for the future of scanning probe microscopy in the study of membrane proteins in transporting epithelia is the development of the scanning ion conductance microscope. Various versions of scanning ion conductance microscopes are presently being developed for biological applications by a number of groups (38, 39, 44). In essence, the instrument is a modified version of the AFM. In place of a conventional probe, however, is a glass micropipette that senses force and ion currents as it scans over the surface. In one version of the microscope (38, 39) it is not necessary for the micopipette/probe to make physical contact with the substrate to produce a conductance map. This feature should allow the generation of maps of conducting epithelial cells that add a new dimension to scanning probe microscopy.


    ACKNOWLEDGEMENTS

We thank the staff of Digital Instruments in Mannheim for valuable technical support. The paper is dedicated to our friend and teacher, Prof. Gerhard Giebisch, Dept. of Cellular and Molecular Physiology, Yale University, for his longstanding support of and interest in our work.


    FOOTNOTES

H. Oberleithner was supported by Deutsche Forschungsgemeinschaft OB 63/8-1 and by the Interdisziplinäre Zentrum fuer Klinische Forschung (IZKF), Universität Münster.

Address for reprint requests and other correspondence: R. M. Henderson, Dept. of Pharmacology, Univ. of Cambridge, Tennis Court Rd., Cambridge, CB2 1QJ, UK (E-mail: rmh1003{at}cam.ac.uk).


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
OBTAINING IMAGES AND FORCE...
THE AFM AND INTACT...
IMAGING SUBCELLULAR MEMBRANE...
ARTIFICIAL LIPID MEMBRANES AND...
ISOLATED MEMBRANE PROTEINS
SPECIAL TREATMENT OF TIPS
THE SCANNING ION CONDUCTANCE...
REFERENCES

1.   A-Hassan, E, Heinz WF, Antonik MD, D'Costa NP, Nageswaran S, Schoenenberger CA, and Hoh JH. Relative microelastic mapping of living cells by atomic force microscopy. Biophys J 74: 1564-1578, 1998[Abstract/Free Full Text].

2.   Al-Awqati, Q. Regulation of ion channels by ABC transporters that secrete ATP. Science 269: 805-806, 1995[ISI][Medline].

3.   Allen, S, Chen XY, Davies J, Davies MC, Dawkes AC, Edwards JC, Roberts CJ, Sefton J, Tendler SJB, and Williams PM. Detection of antigen-antibody binding events with the atomic force microscope. Biochemistry 36: 7457-7463, 1997[ISI][Medline].

4.   Allen, S, Davies J, Dawkes AC, Davies MC, Edwards JC, Parker MC, Roberts CJ, Sefton J, Tendler SJB, and Williams PM. In situ observation of streptavidin-biotin binding on an immunoassay well surface using an atomic force microscope. FEBS Lett 390: 161-164, 1996[ISI][Medline].

5.   Argaman, M, Golan R, Thomson NH, and Hansma HG. Phase imaging of moving DNA molecules and DNA molecules replicated in the atomic force microscope. Nucleic Acids Res 25: 4379-4384, 1997[Abstract/Free Full Text].

6.   Beckmann, M, Nollert P, and Kolb HA. Manipulation and molecular resolution of a phosphatidylcholine- supported planar bilayer by atomic force microscopy. J Membr Biol 161: 227-233, 1998[ISI][Medline].

7.   Binnig, G, Quate CF, and Gerber C. Atomic force microscope. Phys Rev Lett 56: 930-933, 1986[ISI][Medline].

8.   Bottomley, LA, Coury JE, and First PN. Scanning probe microscopy. Anal Chem 68: 185R-230R, 1996.

9.   Bourdieu, L, Ronsin O, and Chatenay D. Molecular positional order in Langmuir-Blodgett films by atomic force microscopy. Science 259: 798-801, 1993[ISI].

10.   Bustamante, JO, Liepins A, Prendergast RA, Hanover JA, and Oberleithner H. Patch clamp and atomic force microscopy demonstrate TATA-binding protein (TBP) interactions with the nuclear pore complex. J Membr Biol 146: 263-272, 1995[ISI][Medline].

11.   Czajkowsky, DM, and Shao ZF. Submolecular resolution of single macromolecules with atomic force microscopy. FEBS Lett 430: 51-54, 1998[ISI][Medline].

12.   Dai, HJ, Hafner JH, Rinzler AG, Colbert DT, and Smalley RE. Nanotubes as nanoprobes in scanning probe microscopy. Nature 384: 147-150, 1996[ISI].

13.   Danker, T, Mazzanti M, Tonini R, Rakowska A, and Oberleithner H. Using atomic force microscopy to investigate patch-clamped nuclear membrane. Cell Biol Int 21: 747-757, 1997[ISI][Medline].

14.   Ehrenhöfer, U, Rakowska A, Schneider SW, Schwab A, and Oberleithner H. The atomic force microscope detects ATP-sensitive protein clusters in the plasma membrane of transformed MDCK cells. Cell Biol Int 21: 737-746, 1997[ISI][Medline].

15.   Ellis, DJ, Berge T, Edwardson JM, and Henderson RM. Investigation of protein partnerships using atomic force microscopy. Microsc Res Tech 44: 368-377, 1999[ISI][Medline].

16.   Engel, A, Schoenenberger CA, and Müller DJ. High resolution imaging of native biological sample surfaces using scanning probe microscopy. Curr Opin Struct Biol 7: 279-284, 1997[ISI][Medline].

17.   Florin, E-L, Moy VT, and Gaub HE. Adhesion forces between individual ligand-receptor pairs. Science 264: 415-417, 1994[ISI][Medline].

18.   Florin, EL, Rief M, Lehmann H, Ludwig M, Dornmair C, Moy VT, and Gaub HE. Sensing specific molecular interactions with the atomic force microscope. Biosens Bioelectron 10: 895-901, 1995[ISI].

19.   Folprecht, G, Schneider S, and Oberleithner H. Aldosterone activates the nuclear pore transporter in cultured kidney cells imaged with atomic force microscopy. Pflügers Arch 432: 831-838, 1996[ISI][Medline].

20.   Goldie, KN, Panté N, Engel A, and Aebi U. Exploring native nuclear-pore complex structure and conformation by scanning force microscopy in physiological buffers. J Vacuum Sci Technol 12: 1482-1485, 1994[ISI].

21.   Grandbois, M, Clausen-Schaumann H, and Gaub H. Atomic force microscope imaging of phospholipid bilayer degradation by phospholipase A2. Biophys J 74: 2398-2404, 1998[Abstract/Free Full Text].

22.   Hansma, HG, Kim KJ, Laney DE, Garcia RA, Argaman M, Allen MJ, and Parsons SM. Properties of biomolecules measured from atomic force microscope images: a review. J Struct Biol 119: 99-108, 1997[ISI][Medline].

23.   Hansma, HG, and Pietrasanta L. Atomic force microscopy and other scanning probe microscopies. Curr Opin Chem Biol 2: 579-584, 1998[ISI][Medline].

24.   Hansma, PK, Cleveland JP, Radmacher M, Walters DA, Hillner PE, Bezanilla M, Fritz M, Vie D, Hansma HG, Prater CB, Massie J, Fukunaga L, Gurley J, and Elings V. Tapping mode atomic force microscopy in liquids. Appl Phys Lett 64: 1738-1740, 1994[ISI].

25.   Hansma, PK, Drake B, Grigg D, Prater CB, Yashar F, Gurley G, Eling SV, Feinstein S, and Lal R. A new, optical-lever based atomic-force microscope. J Appl Physiol 76: 796-799, 1994[ISI].

26.   Hansma, PK, Elings VB, Marti O, and Bracker CE. Scanning tunneling microscopy and atomic force microscopy: application to biology and technology. Science 242: 209-216, 1988[ISI][Medline].

27.   Heginbotham, L, Odessey E, and Miller C. Tetrameric stoichiometry of a prokaryotic K+ channel. Biochemistry 36: 10335-10342, 1997[ISI][Medline].

28.   Henderson, E, Haydon PG, and Sakaguchi DS. Actin filament dynamics in living glial cells imaged by atomic force microscopy. Science 257: 1944-1946, 1992[ISI][Medline].

29.   Henderson, RM. High resolution imaging of biological macromolecules using the atomic force microscope. Exp Nephrol 5: 453-456, 1997[ISI][Medline].

30.   Henderson, RM, Schneider S, Li Q, Hornby D, White SJ, and Oberleithner H. Imaging ROMK1 inwardly-rectifying ATP-sensitive K+ channel using atomic force microscopy. Proc Natl Acad Sci USA 93: 8756-8760, 1996[Abstract/Free Full Text].

31.   Hinterdorfer, P, Baumgartner W, Gruber HJ, Schilcher K, and Schindler H. Detection and localization of individual antibody-antigen recognition events by atomic force microscopy. Proc Natl Acad Sci USA 93: 3477-3481, 1996[Abstract/Free Full Text].

32.   Ho, K, Nichols CG, Lederer WJ, Lytton J, Vassilev PM, Kanazirska MV, and Hebert SC. Cloning and expression of an inwardly rectifying ATP-regulated potassium channel. Nature 362: 31-38, 1993[ISI][Medline].

33.   Hoh, JH, and Schoenenberger CA. Surface morphology and mechanical properties of MDCK monolayers by atomic force microscopy. J Cell Sci 107: 1105-1114, 1994[Abstract/Free Full Text].

34.   Ikai, A. STM and AFM of bio/organic molecules and structures. Surface Sci Rep 26: 263-332, 1996[ISI].

35.   Kasas, S, Thomson NH, Smith BL, Hansma PK, Miklossy J, and Hansma HG. Biological applications of the AFM: From single molecules to organs. Int J Imaging Systems Technol 8: 151-161, 1997[ISI].

36.   Kato, K, Makino Y, Kishimoto T, Yamauchi J, Kato S, Muramatsu M, and Tamura T. Multimerization of the mouse TATA-binding protein (TBP) driven by its C-terminal conserved domain. Nucleic Acids Res 22: 1179-1185, 1994[Abstract].

37.   Knapp, HF, Wiegräbe W, Heim M, Eschrich R, and Guckenberger R. Atomic force microscope measurements and manipulation of Langmuir-Blodgett films with modified tips. Biophys J 69: 708-715, 1995[Abstract].

38.   Korchev, YE, Bashford CL, Milovanovic M, Vodyanoy I, and Lab MJ. Scanning ion conductance microscopy of living cells. Biophys J 73: 653-658, 1997[Abstract].

39.   Korchev, YE, Milovanovic M, Bashford CL, Bennett DC, Sviderskaya EV, Vodyanoy I, and Lab MJ. Specialized scanning ion-conductance microscope for imaging of living cells. J Microsc 188: 17-23, 1997[ISI][Medline].

40.   Lal, R, and John SA. Biological applications of atomic force microscopy. Am J Physiol Cell Physiol 266: C1-C21, 1994[Abstract/Free Full Text].

41.   Lärmer, J, Schneider SW, Danker T, Schwab A, and Oberleithner H. Imaging excised apical plasma membrane patches of MDCK cells in physiological conditions with atomic force microscopy. Pflügers Arch 434: 254-260, 1997[ISI][Medline].

42.   Lee, MG, Wigley WC, Zeng WZ, Noel LE, Marino CR, Thomas PJ, and Muallem S. Regulation of Cl-/HCO-3 exchange by cystic fibrosis transmembrane conductance regulator expressed in NIH 3T3 and HEK 293 cells. J Biol Chem 274: 3414-3421, 1999[Abstract/Free Full Text].

43.   Le Grimellec, C, Lesniewska E, Cachia C, Schreiber JP, De Fornel F, and Goudonnet JP. Imaging of the membrane surface of MDCK cells by atomic force microscopy. Biophys J 67: 36-41, 1994[Abstract].

44.   Le Grimellec, C, Lesniewska E, Giocondi MC, Finot E, and Goudonnet JP. Simultaneous imaging of the surface and the submembraneous cytoskeleton in living cells by tapping mode atomic force microscopy. C R Acad Sci III 320: 637-643, 1997[ISI][Medline].

45.   Le Grimellec, C, Lesniewska E, Giocondi MC, Finot E, Vié V, and Goudonnet JP. Imaging of the surface of living cells by low-force contact-mode atomic force microscopy. Biophys J 75: 695-703, 1998[Abstract/Free Full Text].

46.   Lesniewska, E, Giocondi MC, Vi V, Finot E, Goudonnet JP, and Le Grimellec C. Atomic force microscopy of renal cells: Limits and prospects. Kidney Int Suppl 65: S42-S48, 1998[Medline].

47.   Li, HL, Sui HX, Ghanshani S, Lee S, Walian PJ, Wu CL, Chandy KG, and Jap BK. Two-dimensional crystallization and projection structure of KcsA potassium channel. J Mol Biol 282: 211-216, 1998[ISI][Medline].

48.   Li, Q, Cope G, Hornby D, and White S. Immunocytochemical location of the potassium channel ROMK1 in rat kidney cortex (Abstract). J Physiol (Lond) 489: 93P-94P, 1995.

49.   McTigue, MA, Williams DR, and Tainer JA. Crystal structures of a schistosomal drug and vaccine target: glutathione S-transferase from Schistosoma japonica and its complex with the leading antischistosomal drug praziquantel. J Mol Biol 245: 21-27, 1995.

50.   Morris, VJ. Biological applications of scanning probe microscopes. Prog Biophys Mol Biol 61: 131-185, 1994[ISI][Medline].

51.   Müller, DJ, Engel A, and Amrein M. Preparation techniques for the observation of native biological systems with the atomic force microscope. Biosens Bioelectron 12: 867-877, 1997[ISI].

52.   Müller, DJ, Schoenenberger CA, Schabert F, and Engel A. Structural changes in native membrane proteins monitored at subnanometer resolution with the atomic force microscope: a review. J Struct Biol 119: 149-157, 1997[ISI][Medline].

53.   Neagu, C, Van der Werf KO, Putman CAJ, Kraan YM, De Grooth BG, Van Hulst NF, and Greve J. Analysis of immunolabeled cells by atomic force microscopy, optical microscopy, and flow cytometry. J Struct Biol 112: 32-40, 1994[ISI][Medline].

54.   Oberleithner, H, Brinckmann E, Giebisch G, and Geibel J. Visualising life on biomembranes by atomic force microscopy. Kidney Int 48: 795-801, 1995.

55.   Oberleithner, H, Brinckmann E, Schwab A, and Krohne G. Imaging nuclear pores of aldosterone-sensitive kidney cells by atomic force microscopy. Proc Natl Acad Sci USA 91: 9784-9788, 1994[Abstract/Free Full Text].

56.   Oberleithner, H, Geibel J, Guggino W, Henderson RM, Hunter M, Schneider SW, Schwab A, and Wang W. Life on biomembranes viewed with the atomic force microscope. Wien Klin Wochenschr 109: 419-423, 1997[ISI][Medline].

57.   Oberleithner, H, Giebisch G, and Geibel J. Imaging the lamellipodium of migrating epithelial cells in vivo by atomic force microscopy. Pflügers Arch 425: 506-510, 1993[ISI][Medline].

58.   Oberleithner, H, Schneider S, and Bustamante JO. Atomic force microscopy visualizes ATP-dependent dissociation of multimeric TATA-binding protein before translocation into the cell nucleus. Pflügers Arch 432: 839-844, 1996[ISI][Medline].

59.   Oberleithner, H, Schneider S, Lärmer J, and Henderson RM. Viewing the renal epithelium with the atomic force microscope. Kidney Blood Press Res 19: 142-147, 1996[ISI][Medline].

60.   Oberleithner, H, Schneider SW, and Henderson RM. Structural activity of a cloned potassium channel (ROMK1) monitored with the atomic force microscope: the "molecular-sandwich" technique. Proc Natl Acad Sci USA 94: 14144-14149, 1997[Abstract/Free Full Text].

61.   Oberleithner, H, Schwab A, Wang W, Giebisch G, Hume F, and Geibel J. Living renal epithelial cells imaged by atomic force microscopy. Nephron 66: 8-13, 1994[ISI][Medline].

62.   Panté, N, and Aebi U. Molecular dissection of the nuclear pore complex. Crit Rev Biochem Mol Biol 31: 153-199, 1996[Abstract].

63.   Perez-Terzic, C, Pyle J, Jaconi M, Stehno-Bittel L, and Clapham DE. Conformational states of the nuclear pore complex induced by depletion of nuclear Ca2+ stores. Science 273: 1875-1877, 1996[Abstract/Free Full Text].

64.   Proksch, R, Lal R, Hansma PK, Morse D, and Stucky G. Imaging the internal and external pore structure of membranes in fluid: tapping mode scanning ion conductance microscopy. Biophys J 71: 2155-2157, 1996[Abstract].

65.   Putman, CAJ, De Grooth BG, Hansma PK, Van Hulst NF, and Greve J. Immunogold labels: cell-surface markers in atomic force microscopy. Ultramicroscopy 48: 177-182, 1993[ISI].

66.   Putman, CAJ, Van der Werf KO, De Grooth BG, Van Hulst NF, and Greve J. Tapping mode atomic force microscopy in liquid. Appl Phys Lett 64: 2454-2456, 1994[ISI].

67.   Radmacher, M, Fritz M, Hansma HG, and Hansma PK. Direct observation of enzyme activity with the atomic force microscope. Science 265: 1577-1579, 1994[ISI][Medline].

68.   Radmacher, M, Fritz M, Kacher CM, Cleveland JP, and Hansma PK. Measuring the viscoelastic properties of human platelets with the atomic force microscope. Biophys J 70: 556-567, 1996[Abstract].

69.   Radmacher, M, Tillman RW, Fritz M, and Gaub HE. From molecules to cells: imaging soft samples with the atomic force microscope. Science 257: 1900-1905, 1992[ISI][Medline].

70.   Rakowska, A, Danker T, Schneider SW, and Oberleithner H. ATP-induced shape change of nuclear pores visualized with the atomic force microscope. J Membr Biol 163: 129-136, 1998[ISI][Medline].

71.   Reichelt, R, Holzenburg A, Buhle EL, Jarnik M, Engel A, and Aebi U. Correlation between structure and mass-distribution of the nuclear-pore complex and of distinct pore complex components. J Cell Biol 110: 883-894, 1990[Abstract].

72.   Schaus, SS, and Henderson ER. Cell viability and probe-cell membrane interactions of XR1 glial cells imaged by atomic force microscopy. Biophys J 73: 1205-1214, 1997[Abstract].

73.   Schneider, S, Folprecht G, Krohne G, and Oberleithner H. Immunolocalization of lamins and nuclear pore complex proteins by atomic force microscope. Pflügers Arch 430: 795-801, 1995[ISI][Medline].

74.   Schneider, SW, Egan ME, Jena BP, Guggino WB, Oberleithner H, and Geibel JP. Continuous extracellular ATP measurement on living human lung epithelial cells using atomic force microscopy. Pflügers Arch 433: 1-57, 1997.

75.   Schneider, SW, Lärmer J, Henderson RM, and Oberleithner H. Molecular weights of individual proteins correlate with molecular volumes measured by atomic force microscopy. Pflügers Arch 435: 362-367, 1998[ISI][Medline].

76.   Schneider, SW, Yano Y, Sumpio BE, Jena BP, Geibel JP, Gekle M, and Oberleithner H. Rapid aldosterone-induced cell volume increase of endothelial cells measured by the atomic force microscope. Cell Biol Int 21: 759-768, 1997[ISI][Medline].

77.   Schoenenberger, CA, and Hoh JH. Slow cellular dynamics in MDCK and R5 cells monitored by time-lapse atomic force microscopy. Biophys J 67: 929-936, 1994[Abstract].

78.   Shao, ZF, Yang J, and Somlyo AP. Biological atomic force microscopy: from microns to nanometers and beyond. Annu Rev Cell Biol 11: 241-265, 1995[ISI][Medline].

79.   Smith, PR, Bradford AL, Schneider S, Benos DJ, and Geibel JP. Localization of amiloride-sensitive sodium channels in A6 cells by atomic force microscopy. Am J Physiol Cell Physiol 272: C1295-C1298, 1997[Abstract/Free Full Text].

80.   Thimonier, J, Montixi C, Chauvin JP, He HT, Rocca-Serra J, and Barbet J. Thy-1 immunolabeled thymocyte microdomains studies with the atomic force microscope and the electron microscope. Biophys J 73: 1627-1632, 1997[Abstract].

81.   Vinckier, A, and Semenza G. Measuring elasticity of biological materials by atomic force microscopy. FEBS Lett 430: 12-16, 1998[ISI][Medline].

82.   Walz, T, Tittmann P, Fuchs KH, Müller DJ, Smith BL, Agre P, Gross H, and Engel A. Surface topographies at subnanometer-resolution reveal asymmetry and sidedness of aquaporin-1. J Mol Biol 264: 907-918, 1996[ISI][Medline].

83.   Weisenhorn, AL, Schmitt FJ, Knoll W, and Hansma PK. Streptavidin binding observed with an atomic force microscope. Ultramicroscopy 42-44: 1125-1132, 1992.

84.   Willemsen, OH, Snel MME, Van der Werf KO, De Grooth BG, Greve J, Hinterdorfer P, Gruber HJ, Schindler H, Van Kooyk Y, and Figdor CG. Simultaneous height and adhesion imaging of antibody-antigen interactions by atomic force microscopy. Biophys J 75: 2220-2228, 1998[Abstract/Free Full Text].

85.   Zasadzinski, JAN, Helm CA, Longo ML, Weisenhorn AL, Gould SAC, and Hansma PK. Atomic force microscopy of hydrated phosphatidylethanolamine bilayers. Biophys J 59: 755-760, 1991[Abstract].


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