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
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ABSTRACT |
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
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INTRODUCTION |
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).
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OBTAINING IMAGES AND FORCE DATA WITH THE AFM |
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.
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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.
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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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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.
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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)].
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THE AFM AND INTACT RENAL CELLS |
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.
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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).]
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IMAGING SUBCELLULAR MEMBRANE STRUCTURES |
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).
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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.
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ARTIFICIAL LIPID MEMBRANES AND RECONSTITUTION OF MEMBRANE PROTEINS |
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 |
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 |
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 |
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).
 |
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