MICROSCOPY RESEARCH AND TECHNIQUE 47:3–17 (1999)
Visualising the Actin Cytoskeleton
J.-VICTOR SMALL,1* KLEMENS ROTTNER,1 PENELOPE HAHNE,1 AND KURT I. ANDERSON2
1Institute
2Marie
of Molecular Biology, Austrian Academy of Sciences, Salzburg, Austria
Curie Research Institute, The Chart, Oxted, Surrey, England
KEY WORDS
cytoskeleton dynamics; digital microscopy; GFP; actin
ABSTRACT
The actin cytoskeleton is a dynamic filamentous network whose formation and
remodeling underlies the fundamental processes of cell motility and shape determination. To serve
these roles, different compartments of the actin cytoskeleton engage in forming specific coupling
sites between neighbouring cells and with the underlying matrix, which themselves serve signal
transducing functions. In this review, we focus on methods used to visualise the actin cytoskeleton
and its dynamics, embracing the use of proteins tagged with conventional fluorophores and green
fluorescent protein. Included also is a comparison of cooled CCD technology, confocal and 2-photon
fluorescence microscopy of living and fixed cells, as well as a critique of current procedures for
electron microscopy. Microsc. Res. Tech. 47:3–17, 1999. r 1999 Wiley-Liss, Inc.
INTRODUCTION
As a major, ubiquitous protein in all metazoan cells,
actin serves central roles in shape determination, cytokinesis, and cell motility, as well as in the establishment of cell–cell and cell–matrix interactions (Amos
and Amos, 1991; Bray, 1992). A notable property of
living cells is their ability to regionally control the
polymerisation and supramolecular organisation of actin filaments, involving the engagement of a broad
spectrum of actin binding proteins and resulting in the
formation of different structural subcompartments, each
with a defined function (e.g., Small et al., 1998, 1999).
In epithelial cells, a circumferential band of actin
filaments provides the structural support for cell–cell
junctions; in motile cells, such as leukocytes, actin
filaments form the meshed framework of the protruding lamellipodia and linear filopodia and immobile
fibroblasts are anchored flat to the underlying matrix
via transmembrane coupling to linear bundles of actin
filaments, or ‘‘stress fibres.’’ In response to external
signals, dramatic changes in cell shape and motility can
be effected and all involve the controlled and dynamic
reorganisation of the actin cytoskeleton.
Discoveries in this decade (Hall, 1998) have revealed
that the Rho family of small GTPases play a central role
in the pathways that signal the formation of the
different actin subcompartments (Fig. 1): lamellipodia
are induced by Rac, filopodia by Cdc42, and stress fibres
by Rho. Current interest now focuses on other members
of these pathways and on the mechanisms by which
they are engaged to effect the polymerisation and
regional organisation of actin filaments. In these studies the living cell as a test tube is playing an essential
role, requiring the use of techniques with which the
dynamic reorganisations of the actin cytoskeleton can
be readily manipulated, recorded, and related to function. Our intention here is to review current procedures
for the visualisation of the actin cytoskeleton in both
fixed and living cells. Advances in microscope and video
camera technology now offer interesting options and in
describing the different options available, we will aim
to give a frank assessment of their advantages and
r 1999 WILEY-LISS, INC.
disadvantages. While the comments here will be restricted to the actin cytoskeleton, we hope that they
may also be of interest to those studying cytoskeleton
dynamics in general.
IMMUNOCYTOCHEMISTRY
For analysis of the subcellular localisation of putative
components of the actin cytoskeleton, indirect immunofluorescence microscopy continues to be a valuable
approach. The labelling of the cytoskeleton with antibodies and phalloidin necessitates steps of chemical fixation and membrane permeabilisation. These two steps
may be combined, or permeabilisation performed after
fixation.
We have previously described in detail the fixation
and permeabilisation options available and the compromises that have to be made in choosing conditions
compatible with good antibody labelling and reasonable
structural preservation (Herzog et al., 1994; Mies et al.,
1998). Since most antibodies show variable reactivities
to cells fixed in different ways, the individual antibody
in use dictates the fixation protocol, requiring test
fixations to optimise the result. As far as the structure
of the actin cytoskeleton is concerned, the preservation
of actin stress fibre bundles is relatively insensitive to
the fixation protocol. This is not the case, however, for
the looser organisations of actin filaments in the body of
the cytoplasm or for the peripheral lamellipodia and
filopodia. In weakly fixed cells, the latter become distorted or even washed away during the labelling procedure.
When only actin filaments need to be visualised,
fluorescent phalloidin (Wulf et al., 1979) is the probe of
choice since this can be used after strong fixation. In
our hands the best fixation for phalloidin labelling
Contract grant sponsors: the Austrian Science Research Council, the Austrian
National Bank, and the Seegen Stiftung of the Austrian Academy of Sciences.
*Correspondence to: Institute of Molecular Biology, Austrian Academy of
Sciences, A-5020 Salzburg, Billrothstrasse 11, Austria. E-mail: jvsmall@imb.
oeaw.ac.at
Received 4 February 1999; accepted in revised form 5 May 1999.
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J.-V. SMALL ET AL.
Fig. 1. Subcompartments in the actin cytoskeleton of a chick heart fibroblast, labelled with
rhodamine-conjugated phalloidin. Abbreviations: sf, stress fibre; lam, lamellipodium; ms/fil microspike/
filopodium; am, actin meshwork; arc, dorsal arc. Image was recorded on negative film.
alone involves brief treatment (1–2 minutes) of cells
with a mixture of glutaraldehyde and Triton X-100
(0.25–0.5% glutaraldehyde; 0.25–0.5% Triton), followed
by 1% glutaraldehyde for 10 minutes (Rinnerthaler et
al., 1988). The buffer used (see Herzog et al., 1994) has a
pH ⬃6.1 and includes MgCl2 and EGTA to stabilise
actin filaments. These particular conditions were tailored from electron microscope studies and yield excellent ultrastructural preservation of the actin cytoskeleton (Small, 1988). To eliminate autofluorescence from
free aldehyde groups, brief treatment with sodium
borohydride is required before phalloidin labelling.
Phalloidin is supplied as a dried residue that is best
dissolved in methanol (0.1 mg/ml) and stored at ⫺20°C,
under which conditions it is stable for many months to
years, depending on the fluorescent derivative. For very
long storage, aliquots from the methanol stock can be
vacuum-dried and kept at ⫺20°C. The best fluorescent
phalloidin probes on the market at present are rhodamine phalloidin (Sigma, St. Louis, MO, or Molecular
Probes, Eugene, OR) and the new Alexa 488 (green) and
Alexa 568 (red) phalloidins (Molecular Probes). A Cy-3
phalloidin, produced by Prof. H. Faulstich (Heidelberg)
is also an excellent probe, superior to rhodamine phalloidin. Since phalloidin binds only filamentous actin,
measurements of phalloidin intensity through digital
imaging can be used to quantitate relative filament
densities within and between different actin cytoskeleton subcompartments. Such measurements also serve
as a guide to the degree of preservation of the actin
cytoskeleton under different fixation conditions (see
e.g., Small et al., 1995).
The advantage of phalloidin is that it can be applied
to cells in an aqueous buffer in a simple, one-step
procedure. Normally, a brief (30-minute) incubation
suffices, but if the labelling is weak it can be continued
overnight at 4°C, sometimes with surprisingly improved results. When combined with antibody labelling, phalloidin can be included in any step of the
procedure. And since phalloidin has a stabilising effect
on actin filaments, it can usefully be applied early
during antibody labelling, especially to help stabilise
cells that are weakly fixed to satisfy the antibody
requirements.
We will not attempt to review work dealing with the
production and use of actin antibodies, since useful
commercial antibodies are mainly directed against
specific actin isotypes (e.g., Tsukada et al., 1987; Skalli
et al., 1986; Gimona et al., 1994). Since these are
generally monoclonals, they are normally less suitable
than phalloidin as secondary actin markers in combination with other monoclonal antibodies directed against
cytoskeleton components. Nevertheless, such antibodies are important tools in studies of actin isoform
VISUALISING THE ACTIN CYTOSKELETON
distribution, but care must be taken to screen different
fixation procedures to obtain consistent results. For
example, we have found that batches of the monoclonal
beta actin antibody, first characterised by us (Gimona
et al., 1994), can show restricted, mainly lamellipodia
staining with aldehyde fixation, but labelling of the
entire cytoskeleton after fixation in methanol (Mies et
al., 1998). It can be anticipated that GFP analogues of
actin isoforms (see further below) will supersede isoform-specific antibodies in studies of isoform distribution, at least in cultured cells.
OBSERVATION OF ACTIN FILAMENT
DYNAMICS IN LIVING CELLS
Techniques for following the reorganisations of actin
filaments in living cells are based on the method of
‘‘fluorescent analog cytochemistry’’ (Wang, 1989), entailing the conjugation of proteins with fluorescent probes
and their introduction into the cytoplasm by microinjection, electroporation, or scrape loading (Wang, 1994;
Fig. 2a,c). The detection of fluorescent protein analogs
in living cells relies on the use of important developments in video microscopy, involving the employment of
high sensitivity, cooled CCD cameras together with
controlled shutter devices to minimise radiation damage (Inoue and Spring, 1997). The characteristics of the
imaging systems will be discussed in more detail in the
section Imaging Living and Fixed Cells.
The first studies with fluorescent conjugates of cytoskeleton proteins showed that the labelled proteins
localised correctly to the sites occupied by the endogenous protein, determined independently by immunofluorescence microscopy (reviewed in Jockusch et al.,
1986; Wang and Sanders, 1990). With actin, it could be
shown in fibroblasts that uptake and turnover was
highest in the peripheral lamellipodia and lowest in the
stress fibres (Kreis et al., 1982; Glacy, 1983; Turnacioglu et al., 1998; Wang, 1984). Special interest has
focused on the dynamics of actin filament turnover in
lamellipodia, since their protrusion is essential for cell
motility. By combining fluorescent actin microinjection
with spot photobleaching using a focused laser beam,
Wang (1985) was able to demonstrate a rearward flow
of actin in lamellipodia. The same rearward flow was
demonstrated by two further techniques that will no
doubt come into more general use: activation of caged
fluorescence (Theriot and Mitchison, 1991; Theriot et
al., 1992) and speckled labelling (Waterman-Storer et
al., 1998).
Caged fluorescent probes are very useful because
they provide a better alternative to photobleaching for
following turnover events. In essence, the technique
involves the coupling of a protein to a caged probe and
its injection into cells. An intense beam of light of the
appropriate wavelength, focused to a spot or line, is
then used to uncage the probe, causing it to fluoresce
and the dynamics of the labelled protein is then determined from the fate of the fluorescent zone. However,
the use of caged probes in studying cytoskeleton dynamics, although having already produced dramatic results
(Mitchison, 1989; Theriot and Mitchison, 1991; Theriot
et al., 1992) is not yet a routine technique. Mitchison et
al. (1994) described the synthesis of caged probes and
their conjugation to proteins, but also indicated the
need for improvements. Some caged probes are also
5
available commercially, but we have encountered unexpected problems in uncaging actin conjugates in a
cellular environment as compared to free in solution
(Anderson, unpublished results). The development of
new probes is apparently necessary to make this technique more widely applicable.
The speckled technique is based on the observation
that the injection of low amounts of fluorescent tubulin
or actin did not give simply a lower overall density of
fluorescent label in the injected cells, but instead a
spotty or ‘‘speckled’’ pattern (Waterman-Storer et al.,
1998). The fact that the speckles flowed in a way
predicted by other techniques for the turnover of tubulin and actin suggested that the punctate label reflected
functional incorporation into cytoskeletal filaments.
This method offers an interesting alternative to activated fluorescence in following the dynamics of actin or
tubulin. In particular, it might be usefully combined
with secondary fluorescent probes, complementary to
the speckled label, to relate polymer flow to other
features of cytoskeleton reorganisation and turnover.
Because of the need of the highest working magnifications and high illumination intensities, special attention is required to reduce photobleaching and radiation
damage. For this reason, the addition of the oxygen
scavenging enzyme preparation ‘‘Oxyrase’’ to the growth
medium has been recommended (Waterman-Storer et
al., 1998).
A note about the use of fluorescent phalloidin in
living cells is called for to close this section. Since
phalloidin is such a specific label for actin filaments,
fluorescent analogs were injected into cells as soon as
they were available (Wehland et al., 1977). It was
immediately apparent, however, that phalloidin disrupted actin cytoskeleton organisation which, in retrospect, can be attributed to phalloidin’s inhibition of
actin filament turnover. Nevertheless, it appears that
limited doses of phalloidin can usefully be employed in
some situations to visualise actin in living cells (Wehland and Weber, 1981; Wang, 1987). Sanders and Wang
(1990) employed the injection of short actin filaments,
prelabelled with phalloidin, to demonstrate that the
polymerisation of actin in living cytoplasm is not spontaneous, but a controlled and site-specific phenomenon.
GREEN FLUORESCENT PROTEIN
The possibility to tag previously cloned proteins to
the natural green fluorescent protein (GFP) by molecular genetics (Chalfie et al., 1994) has now added an
important dimension to the general approach of fluorescent analog cytochemistry. Introduction of the engineered c-DNA into cells is achieved either by transfection or microinjection. More recently, a variation of the
GFP-tagging technique has been described involving
the separate transfection of GFP and the desired protein, but with each tagged with a leucine zipper so that
they pair up in the cell (Katz et al., 1998). The generation of GFP in new colours promises the possibility of
following multiple probes in living cells by this technology.
Various studies showing the successful tagging of
GFP to cytoskeletal proteins have appeared (Westphal
et al., 1997; Ludin and Matus, 1998; Fischer et al.,
1998; Ballestrum et al., 1998; Choidas et al., 1998; Pang
et al., 1998) and many more can be expected. We
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J.-V. SMALL ET AL.
Fig. 2. Examples of fluorescent probes introduced into living cells.
a,c: Rhodamine-conjugated and microinjected proteins. b,d: Proteins
tagged with GFP. In a and b the distribution of two focal contact
proteins, paxillin and vinculin, is visualised in the same transfected
and microinjected B16 melanoma cell in separate fluorescent channels. Note identical localisations. The GFP paxillin probe was produced by Marcus Geese, Antonio Sechi, and Jurgen Wehland (Braun-
schweig, Germany) using a cDNA provided by Ravi Salgia and James
Griffin (Boston, MA, USA). c,d: Distribution of actin as seen with
microinjected rhodamine-actin (c, chick fibroblast) and with GFP actin
(d, B16 melanoma cell). The B16 melanoma cells expressing GFP actin
were provided by Christoph Ballestrem, Bernhard Wehrle-Haller, and
Beat A. Imhof (Geneva, Switzerland). All images were recorded on a
back-illuminated, cooled CCD camera (Princeton Instruments).
provide examples here of GFP-actin (Ballestrem et al.,
1998) and GFP-paxillin and compare these with images
of cells injected with fluorescent probes (Figs. 2, 3).
Both the injected vinculin and GFP-paxillin localise
faithfully to the punctate sites of cell substrate adhesion and the actin probes mark the different compartments of the actin cytoskeleton. It remains to be shown,
however, to what extent the turnover of the two types of
Fig. 3. GFP actin as a faithful marker of actin distribution. A living and motile B16 melanoma cell,
expressing GFP actin (a, Ballestrem et al., 1998) was fixed and then labelled with Cy-3 phalloidin (b).
Images obtained on a back-illuminated, cooled CCD camera. Cy-3 phalloidin was provided by Heinz
Faulstich (Heidelberg, Germany).
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J.-V. SMALL ET AL.
probe mimic each other under conditions of prolonged
observation, or in response to manipulations that effect
rapid cytoskeletal changes.
It is notable that Ballestrem et al. (1998) found that
the melanoma cells they employed were sensitive to
overexpression of actin, effected via a viral promoter. To
retain normal cell morphology, the beta actin promoter
was required. In our hands, fibroblasts tried so far were
best transfected using the CMV viral promoter, indicating the need to employ different strategies according to
cell type and to consider the effects that exogenous
expression may exert on cell behaviour.
IMAGING LIVING AND FIXED CELLS USING
COOLED CCD CAMERAS AND LASER
SCANNING SYSTEMS
Digital Imaging
The availability of highly sensitive CCD (charged
coupled device) cameras (Inoue and Spring, 1997) and
laser scanning microscopes (White et al., 1987; Pawley,
1990) now provides digital imaging options, in addition
to classical photography for fluorescence microscopy.
Negative film should still be held in store for routine
work and can deliver the best images of fixed, flat cells,
given intense fluorescent labelling. But digital imaging
offers obvious advantages of data storage and processing, as well as on-line focusing and video capabilities.
We consider here the practical aspects of imaging fixed
and living cells labelled with fluorescent probes. This
will include a brief consideration of the principles of
cooled CCD devices and of confocal and two-photon
laser scanning microscopes.
In fluorescence imaging there is an important practical distinction between dealing with living and fixed
cells. A greater flexibility is allowed in the labelling of
fixed cells to amplify weak fluorescent signals (e.g.,
biotin-avidin systems) and to reduce photobleaching
through additives in the embedding medium. Live cell
imaging is, however, constrained by such factors as
motion within the sample, the amount of fluorescent
probe which can be safely introduced into the cell,
photodamage of the probe, and phototoxic effects on the
cell, requiring more stringent conditions.
The primary strength of the confocal laser scanning
(CLS) microscope lies in the ability to optically section a
specimen. However, its wide availability has led to its
increased use simply as a means of digital microscopy,
i.e., in situations where the depth discrimination and/or
blur reduction is of little significance. Rather than
attempt a detailed technical comparison of CCD and
confocal imaging, we will concentrate on principal
differences of how light from the sample is detected and
the practical implications for viewing cultured cells.
Serial vs. Parallel Illumination
In a CCD camera, a real image of the specimen is
formed on the surface of the chip, which is divided into a
regular array of small units called pixels. Photons
arriving from a given point in the specimen are converted into electrons within each pixel. These electrons
remain stored in the pixel until the chip is read out,
whereby the electrons from each pixel are counted. In
the laser scanning technique, a comparable optical
image of the specimen is never formed (see Sheppard
and Shotton, 1997). Instead, each point in the sample is
illuminated sequentially by a focused and sweeping
laser beam. The light emitted from each specimen point
at the focus of the laser is imaged onto a pin-hole
aperture, which rejects light from all other points in the
specimen. A photomultiplier tube behind the aperture
generates a DC current, whose magnitude is proportional to the number of photons arriving from the
illuminated specimen point.
Thus, in a CCD camera all points in the specimen are
illuminated at once (in parallel), but the corresponding
pixels are read out serially, whereas scanning laser
microscopy relies on both serial illumination and serial
read-out of each point in the specimen. For a CCD
camera it is common to collect an image by exposing a
512 by 512 (262,144) array of pixels for 1 second,
whereas 1-second exposure per pixel using the laser
scanning technique would theoretically require approximately 73 hours! The difference in the manner of
illumination dictates different approaches to imaging
with CCD and CLS systems. Thus, to compensate for
the reduced time of illumination per pixel, the laser
scanning technique relies on intense illumination to
produce a burst of fluorescence as the laser sweeps by,
with a dwell time per pixel of several microseconds.
Saturation and Sensitivity
The number of photons which can be emitted from a
diffraction-limited sample volume depends on the number of fluorophores present and the rate of fluorescence
emission per fluorophore. During illumination, the
number of viable fluorophores will be gradually reduced
by photobleaching. As the rate of fluorescence emission
rises, the proportion of fluorophores in triplet states,
long-lasting (10 µs) states in which the molecule is
unable to fluoresce, will increase, further reducing the
number of photons which can be emitted under intense
illumination (Wells et al., 1990). Thus, at high laser
intensity fluorescence emission becomes saturated and
the signal can only be increased by adding several weak
scans together.
Saturation is generally not such a limiting factor in
cooled CCD microscopy; increased fluorescence signal is
easily obtained by increasing the length of the exposure, with exposures of several seconds common. Should
the observation of rapid events require short exposures,
the level of illumination can be increased, although this
is usually done only as a last resort in order to minimise
phototoxic effects. Cooled CCD cameras, especially the
thinned back-illuminated variety, are generally more
sensitive due to higher quantum efficiently (QE), the
efficiency with which photons arriving from the sample
are converted into countable electrons. The QE of
cooled, back-illuminated CCDs is at least 75% in the
green and red, meaning that for every 100 photons
which arrive at the chip, 75 are converted into countable electrons. In contrast, the QE of standard photomultiplier tubes (PMT) used for LSM is at best 23% (often
less), which places a significant limit on the detection of
low signals.
Optical Sectioning
The relative thinness of most cultured cells (ⱖ5 µ)
makes them inappropriate for optical sectioning, the
process whereby an image is collected from a single
focal plane in the specimen. However, as shown in
VISUALISING THE ACTIN CYTOSKELETON
9
Fig. 4. Images of living and locomoting keratocytes injected with
TAMRA (tetramethylrhodamine) conjugated vinculin, obtained with a
cooled CCD-camera (a) and a CLSM (b). Vinculin is incorporated at
sites of contact with the substrate. Cells are moving from the top to
bottom in the field, with the thick cell bodies (⬃10 µm) trailing behind
broad and thin lamellipodia (0.2–0.3 µm thick). The high background
in the CCD image is caused by the fluorescence of unincorporated
vinculin in the cytoplasm, outside the focal plane of the contacts;
better contrast is achieved in the CLS image because this out-of-focus
fluorescence is not detected.
Figure 4, the ability to reject light from outside the focal
plane remains a powerful advantage of CLSM in reducing background fluorescence in the perinuclear region.
In this example with moving keratocytes, the substrate
contact sites labelled by injecting fluorescent vinculin
are clearly resolved under the cell body in the CLSM,
whereas they are mainly obscured in the CCD image.
Similar results can be obtained using total internal
reflection microscopy (TIRF); however, data collection
is restricted to a region within ⬃150 nm to 1 micron of
the substrate (Axelrod, 1981) and the method is technically demanding.
Optical sectioning can also be achieved using a CCD
camera in conjunction with digital deconvolution, a
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J.-V. SMALL ET AL.
mathematical process which removes out of focus light
from a series of images taken from consecutive specimen planes (Agard and Sedat, 1983). This technique is
limited by the computational time required to generate
the final de-blurred images, from minutes to hours, and
cannot be used when there is insufficient contrast
between the signal from a single focal plane and the
background due to a large volume (e.g., Fig. 4a).
Two-Photon Excitation
Scanning laser microscopy based on two-photon excitation was first demonstrated by Denk et al. (1990). To
understand the technique, some comments on basic
principles are required. Briefly, fluorescence may be
defined as the photon emission which occurs when a
molecule returns from an excited to a ground state. In
single-photon fluorescence, the excited state results
from the absorption of a photon containing the right
amount of energy according to the formula E ⫽ hv,
where h is Planck’s constant and v the reciprocal of the
wavelength. In two-photon fluorescence, the energy
required for excitation is supplied by the simultaneous
absorption of two photons of light of twice the exciting
wavelength and half the required amount of energy (Xu
et al., 1996). This places the energy quanta required for
two-photon excitation of standard fluorophores in the
realm of the infrared, where proteins and nucleic acids
generally do not absorb.
The probability of a fluorophore being struck simultaneously by two photons depends on the local photon
concentration. When a beam of infrared light is focused
to a diffraction limited spot by a microscope objective,
the highest concentration of photons will occur in the
objective’s focal plane. The probability of simultaneous
two-photon absorption will therefore be highest at the
focal point, and vanishingly small elsewhere. Thus the
depth discrimination of the two-photon technique depends on the selective excitation of the sample only in
the focal plane of the objective, whereas depth discrimination in the confocal technique relies on rejection of
out-of-focus light at the detector.
Systems Compared
In Figure 5 we show images of fixed and phalloidinstained keratocytes obtained with a cooled CCD camera
(Fig. 5a), a confocal (Fig. 5b), and a two-photon microscope (Fig. 5c). According to expectations, the resolution
of the two-photon microscope is the lowest due to a
longer wavelength illumination and consequently larger
spot size. In practice, we have found that when excited
to give comparable fluorescent signals, common fluorophores, such as Alexa-488 and GFP, bleach up to 3–5
times more rapidly under two-photon than singlephoton stimulation (D. Drummond and K. Anderson,
unpublished observations). In other words, although
photobleaching outside the focal plane of the objective
is reduced, bleaching in the focal plane proceeds much
more rapidly.
The two-photon system remains advantageous for
the imaging of thick specimens, where light scattering
poses special problems for illumination and detection.
Because scattering and absorption of IR by biological
material is much lower than for visible light, it is
possible to attain intense illumination deep within the
sample (Yuste and Denk, 1995) with reduced photodam-
age compared to the CLSM. Furthermore, in the twophoton system it is certain that all fluorescence returning from the sample was emitted from the same point.
This allows the use of wide-field detection, which
recovers scattered fluorescence lost in confocal detection and can easily boost the signal from deep sections
by 3 to 5 times.
The two-photon system is not well suited for the
standard imaging of thin samples, like cultured cells.
Real-time imaging in this case falls within the scope of
the CLS, as well as the cooled CCD. For specific
purposes, such as monitoring the substrate contacts of
a cell in its thicker regions, CLS has obvious advantages (Fig. 4), but at the expense of sensitivity. Thin
regions of cells are most conveniently recorded with a
cooled CCD camera. One interesting advantage offered
by CLS, however, is the option of parallel interference
reflection microscopy and fluorescence, providing the
possibility to simultaneously compare contact phenomena by two independent techniques. In conclusion,
differences between CLS and CCD systems with regard
to sensitivity and depth discrimination provide complementary approaches for studies of the cytoskeleton.
PRESERVING ACTIN CYTOSKELETON
ULTRASTRUCTURE
General Considerations
The individual filamentous components of the cytoskeleton were originally identified by electron microscope observations of tissue thin sections, cell homogenates, and the purified polymers. The cases of the
muscle cell (actin and myosin filaments), the flagellum
(microtubules), neurons (neuronal intermediate filaments and microtubules), and skin (keratin intermediate filaments) are familiar examples of tissues in which
the filaments were early resolved in thin sections
(Fawcett, 1966). However, it was only through the
application of immunofluorescence microscopy to cultured cells in the mid-1970s that the ubiquitous nature
of the cytoskeleton and its general pervasion of cytoplasmic space was recognised. Since cultured cells are well
suited to investigations by light microscopy, they are
ideal for studies of dynamic processes. In particular,
they offer the opportunity of correlating phenomena
seen in living cells with the underlying structural
organisation. We have already seen examples of such
correlations by fluorescence microscopy; here we survey
methods for visualising cultured cell cytoskeletons in
the electron microscope, the emphasis being on actin.
But first, some comments about choosing conditions
and preserving actin filaments.
For the results at the ultrastructural level to be
meaningful, the preparative methods employed should
minimise structural distortions as well as the loss of
material belonging to the cytoskeleton. We should here
be reminded that the conditions used for visualising the
cytoskeleton in the light microscope cannot necessarily
be extrapolated to the electron microscope. In particular, electron microscope procedures generally require
conditions that quantitatively remove the cell membrane, whereas for immunofluorescence microscopy
only enough membrane must be destroyed to allow
penetration of the antibody probes. Necessity may call
for procedures that work for both immunofluorescence
and electron microscopy, so that direct correlations are
Fig. 5. Images of fixed, fish keratocytes stained with fluorescent
phalloidin (Alexa 488 phalloidin) obtained in a CCD camera, confocal,
and two-photon microscope, as indicated. Confocal and two-photon
images were generated using an MRC1024 confocal microscope (BioRad, Cambridge, MA) equipped with a Tsunami tuneable infrared
laser (Spectra-Physics). For the confocal image, the diameter of the
confocal aperture was set to 2 mm, for the two-photon image the
aperture was opened to nearly full size (6 mm) in order to approximate
the resolution of wide-field detection. Note that it is possible to use
two-photon illumination in conjunction with confocal detection, in
which case the image in (c) would appear identical to (b). However,
confocal detection is far less sensitive than wide-field, requiring
increased fluorescence emission and therefore resulting in increased
photobleaching and phototoxicity.
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J.-V. SMALL ET AL.
possible (e.g., Rinnerthaler et al., 1988; Svitkina et al.,
1995). In other cases, it may be desirable to take the
same cell studied in the light microscope into the
electron microscope (e.g., Heath and Dunn, 1978;
Rinnerthaler et al., 1991; Svitkina et al., 1995) and
where antibody labelling is not a requirement, fewer
restraints are then imposed on the method of primary
fixation. By employing actin-GFP or fluorescent phalloidin as probes it is possible to control for the change of
local filament density in response to extraction and
fixation. This can be done by density-scanning fluorescent images captured in a cooled CCD camera (Small et
al., 1995).
Since we focus on the actin cytoskeleton, some remarks about actin filament preservation are particularly pertinent. Where actin filaments occur in bundles
in cells, decorated and crosslinked by actin-associated
proteins, they are relatively resistant to the rigours of
postfixation, dehydration, and embedding required for
thin-section electron microscopy (Goldman et al., 1976:
Heuser and Kirschner, 1980; Tilney et al., 1998). However, individual filaments or filament networks are
more sensitive to the same manipulation procedures. In
particular, both purified actin filaments and the networks of actin filaments found in lamellipodia are
destroyed by osmium tetroxide (Maupin-Szamier and
Pollard, 1978; Small, 1981) and distorted by dehydration (Small, 1981, 1985). This is not so with microtubules and intermediate filaments. The noted sensitivity
of actin networks explains why little filament order has
been detected in lamellipodia in embedded material
(e.g., Abercrombie et al., 1971; Yamada et al., 1971;
Heath and Dunn, 1978). Just as bundling and crosslinking proteins confer actin with resistance to distortion, a
protective effect is also provided by the decoration of
actin with myosin heads (Ishikawa et al., 1969; Begg et
al., 1978). But such treatment requires that the cells
are extracted and unfixed for myosin decoration, a
procedure that can itself lead to filament loss and the
modification of filament organisation.
In discussing the available options for electron microscopy, we will consider only whole-mount methods in
which whole cytoskeletons of substrate bound cells are
processed for viewing under the electron beam. The
principle is not new and in fact was introduced by
Porter et al. (1945) before the advent of the ultramicrotome, but with whole cells rather than cytoskeletons. In
the following, we consider three procedures that yield
useful images of the cytoskeleton: negative staining,
quick freezing, and critical point drying.
Negative Staining
Negative staining as a technique has contributed
decisively to the resolution of the structure of the actin
filament, to the localisation of associated molecules on
its surface, and to the characterisation of bundled
assemblies of actin filaments (reviewed in Amos and
Amos, 1991; Steinmetz et al., 1997). For isolated actin
filaments, the substructure is preserved in filaments
either absorbed to the support film or suspended in a
thin film of stain (Craig et al., 1980). Because of the
simplicity of the negative staining method (see Bremer
et al., 1998), namely, drying in a heavy metal salt, it
was an obvious first choice for applying to cytoskeletons
(Edds, 1977; Small and Celis, 1978).
Our own procedure for cytoskeletons was developed
from conditions first found suitable for the detergent
skinning of isolated smooth muscle cells (Small, 1977;
Small and Celis, 1978). It was subsequently improved
empirically by screening for modifications yielding the
best ultrastructural preservation in the electron microscope (Höglund et al., 1980; Small, 1981) as well as the
faithful and rapid arrest of cell movement (Small et al.,
1982; Rinnerthaler et al., 1991). The preparative procedure for electron microscopy has been outlined in detail
elsewhere (Small and Herzog, 1994; Small and Sechi,
1998) and is, briefly, as follows. The cells are first
cultured on nickel, silver, or gold grids carrying support
films of either colloidon or formvar. Once the cells have
spread, or after specific treatments, the grids are
processed through the following steps: 1) a brief rinse in
‘‘cytoskeleton buffer’’ (CB: 150 mM NaCl, 5 mM EGTA,
5 mM Mg Cl2, 5 mM glucose, 10 mM MES, pH 6.1) at
room temperature; 2) simultaneous extraction and
fixation in a mixture of Triton X-100 (0.25–0.5%) and
glutaraldehyde (0.25–0.5%) in CB for 1–2 minutes at
RT; 3) fixation in 1–2% glutaraldehyde in CB for at
least 10 minutes, a rinse in CB; 4) transfer to CB
containing 100 µg/ml unconjugated phalloidin (1–24
hours); 5) negative staining in 2% sodium silicotungstate. Phalloidin serves the important function of stabilising actin filaments against distortion during staining
(Small, 1981).
With this general procedure, the organisation of actin
filaments in the lamellipodia of a variety of cells has
been demonstrated (Höglund et al., 1980; Small, 1981;
Karlsson et al., 1984; Claviez et al., 1986; Rinnerthaler
et al., 1991; Small et al., 1995). Extraction with saponin
at 37°C and staining in phosphotungstic acid has also
yielded comparable images of growth cone lamellipodia
(Lewis and Bridgman, 1992). One common observation
that emerges from these studies is that neutral negative stains, such as sodium silicotungstate and phosphotungstic acid, are best suited for contrasting cytoskeletons. Although acidic uranyl acetate or uranyl formate
produce the best contrast images of purified actin
filaments (Craig et al., 1980; Steinmetz et al., 1997),
they are less suited for contrasting actin networks
(Small, 1981; Small and Sechi, 1998). Drying in uranyl
acetate produces a noticeable distortion of actin filaments in lamellipodia, apparently due to a more pronounced collapse of the network than is evident with
the neutral stains. The further, apparently paradoxical,
advantage of the neutral stains is that they produce
less contrast. This turns out to be a distinct advantage
when producing prints for publication.
The advantage of the negative stain method is that it
is simple and fast and has so far produced the most
ordered images of lamellipodium networks. Contrasting is also not restricted to the exposed surface layers,
as with methods involving metal shadowing, and the
stain can penetrate beneath patches of surface membrane and reveal the structure beneath. A disadvantage
with negative staining is that the contrast can vary
from one preparation to another and in different regions of the same grid. It is also only suitable for the
thinner parts of cells, but these can make up large
areas in cells that are naturally flat or well spread.
Until now, filament binding molecules, known to be
present on actin filament arrays from immunofluores-
VISUALISING THE ACTIN CYTOSKELETON
cence microscopy, have not been structurally identified
in cytoskeletons by negative staining.
We have shown that it is possible to film living cells
on electron microscope grids and then process these for
electron microscopy (Rinnerthaler et al., 1991). The
method is tedious, however, and the number of cells
that survive the procedure is low. In more recent work
(Mies, Rottner, and Small, unpublished data), we rediscovered the feasibility of performing light microscopy of
cells on film-coated coverslips and then processing the
same cells for negative staining and electron microscopy. The technique is a variation of that described by
Buckley (1975), who separated cells on the support
films from the coverslips and transferred them to EM
grids. By this means, it should be easier to combine
observations of the dynamics or immunolabelling of
individual cells with the negative staining method.
This particular combination is offered by the critical
point drying procedure, which we discuss next.
Critical Point Drying
Only dried samples can be viewed in the vacuum of
the electron microscope, but simple air-drying leads to
major distortions due to surface tension effects. These
are avoided in the critical point drying method, in
which drying is performed in a temperature-controlled
pressure chamber that allows the transition from liquid
to gas, above the critical point, achieved via a temperature jump. Carbon dioxide is the gas of choice since the
temperature jump can be performed in the convenient
range of around 15–45°C (see Ris, 1985). For this
method, the preparations must be thoroughly dehydrated in acetone or ethanol before exchange with
liquid CO2 can take place.
Initial results with cytoskeletons prepared by the
critical point method were disappointing (reviewed in
Small, 1988), for at least two reasons. First, in regions
known to be rich in actin the filaments were of irregular
contour and thickness and bore no structural resemblance to actin. Second, the aggregation and distortion
of actin filaments precluded a convincing distinction
between actin and intermediate filaments. In a modification of this method, Svitkina et al. (1995) introduced
metal coating after critical point drying, a procedure
that improved contrast, but not filament order. More
recent modifications of different steps in the procedure
have yielded a marked improvement in filament clarity
and order (Svitkina et al., 1995, 1996, 1997). Changes
that have lead to this improvement include posttreatment of glutaraldehyde-fixed cytoskeletons with tannic
acid, followed by uranyl acetate, steps which presumably reduce distortions caused by dehydration (Small,
1985). The thorough elimination of water, prior to
exchange with dried CO2 (Ris, 1985) and the drying of
samples in a horizontal position between lens paper are
also claimed to contribute to the better result.
The authors have further taken advantage of the idea
(Small et al., 1982) of removing actin specifically from
cytoskeletons with an actin depolymerising protein to
reveal structures that are otherwise partially or totally
obscured. In particular, they have been able to reveal
crosslinks between intermediate filaments and microtubules, identified by immunoelectron microscopy as plectin (Svitkina et al., 1996) and laterally registered
arrays of bipolar filaments as components of stress fibre
13
bundles (Verkhovsky et al., 1995). This procedure lends
itself favourably to immunoelectron microscopy with
gold probes, since the gold particles are well contrasted
against the metal-coated structures (Svitkina et al.,
1996, 1997). It is also amenable to correlated light and
electron microscopy, as the cells can be grown on
coverslips.
Limitations of the technique include the requirement
for detergent extraction without simultaneous fixation
to ensure that all surface membrane is removed, as well
as dehydration in organic solvents, which may contribute to distortions. In addition, the use of metal shadowing brings with it the loss of detail on the ventral cell
surface, relevant, for example, to the problem of actin
organisation in substrate contacts. The advantages of
the method, in being relatively simple and providing
striking and informative images of the cytoskeleton,
are, however, clear and it is recommended that more
groups adopt the improved methodology.
Quick Freezing
The quick-freeze deep etch method, championed by
Heuser and successfully applied to tissues, suspended
cells, and isolated molecules, often with dramatic results (e.g., Hirokawa and Heuser, 1981; Heuser, 1983,
1986) is not equally suited for actin cytoskeletons of
substrate-attached cells (reviewed also in Small, 1988).
The method relies on vitreous freezing, achieved either
by rapid immersion of a sample in liquid gas or by
slamming on a polished metal block at ultra-low temperatures. The samples are then transferred to a
precooled block in a vacuum device and the vitreous ice
sublimed at low temperature to expose the upper
layers, which are then coated with platinum. The metal
replica is subsequently floated off, cleaned, and mounted
on a grid for microscopy. Under the most stringent
conditions of freezing, however, the preservation of
actin filament order in stress fibres and lamellipodia of
cultured cell cytoskeletons (Heuser and Kirschner, 1980)
does not match that achieved by either negative staining (Small et al., 1994; Lewis and Bridgman, 1992) or
critical point drying. This is evident from a comparison
of images of fish keratocyte lamellipodia obtained by
the three different methods (Lee et al., 1993; Small et
al., 1994, 1995; Svitkina et al., 1997; Figs. 6, 7).
A primary problem in quick-freezing cell monolayers
is that they must first be rinsed with distilled water to
remove salt, then with 10% methanol to reduce subsequent ice crystal growth, and then blotted to remove as
much solvent as possible to leave a liquid layer thin
enough (10 µm or less) to ensure vitreous freezing.
Glutaraldehyde-fixed actin filaments are not stable in
distilled water (Small, 1985), but the effect may be
minimal in the short times employed in rinsing. However, blotting is a poorly controlled step and can either
lead to local air-drying or to the retention of too much
liquid, leading to ice crystal damage. With these constraints it is not surprising that the yield of successfully
frozen cells is low. In addition, the instrumentation is
expensive, the running costs high, and the method
technically demanding. It also does not offer the possibility of correlated light and electron microscopy of the
same cells.
14
J.-V. SMALL ET AL.
Fig. 6. Electron micrographs of the lamellipodium of fish keratocyte cytoskeletons prepared by
freeze-drying (a) or negative staining (b). Actin filament linearity is lost in the freeze-dried sample.
Reproduced from Small et al. (1994) with permission. Bars ⫽ 0.2 µm.
VISUALISING THE ACTIN CYTOSKELETON
15
Fig. 7. Electron micrograph of same lamellipodium region in a
keratocyte as in Figure 6, but in a sample prepared by the critical point
procedure, according to Svitkina et al. (1995). Note considerable
improvement in preservation as compared to the more laborious
freeze-drying method (Fig. 6a). The micrograph was provided by
Tatyana Svitkina (Madison, WI, USA). Bar ⫽ 0.2 µm.
The Ultrastructural Challenge
Whereas the structure of many molecules that associate with actin has been resolved by rotary shadowing
electron microscopy (e.g., Tyler et al., 1980; Mabuchi
and Wang, 1991) they have still not been identified on
actin filament arrays in cytoskeletons. Techniques to
resolve them still require development. In the meantime, much can be gained from immunoelectron microscopy to localise them in the cytoskeleton, as long as it is
performed in the context of meaningful preservation of
filament order (Svitkina et al., 1997). Efforts in this
direction are worthwhile, since they promise to contribute significantly to the underlying mechanism of cell
motility. To test the credibility of current models of how
actin filament polymerisation is driven at the front of
the lamellipodium, more detailed information on fila-
ment order and of the distribution of proteins implicated in controlling this process is now required.
CONCLUSIONS
Improvements in digital imaging microscopy and, in
particular, the possibility to tag proteins with GFP by
molecular engineering open new avenues for studying
cytoskeleton dynamics in living cells. Notably, video
microscope systems are now becoming user-friendly
enough to be exploited by cell and molecular biologists
alike, and will no doubt adopt the status of standard
equipment where questions about in vivo functions of
molecules are being asked. Advances in this area are
going to entail the use of the cell as a test tube with
multiple probes, since it will be required to relate the
dynamics or localisation of a specific component with
16
J.-V. SMALL ET AL.
that of a known molecule. In this respect, the development of new GFP analogs with different spectral characteristics is bound to play a central role.
Digital microscopy of living cells containing fluorescently conjugated probes allows the quantitation of
protein turnover in different subcellular localisations.
For actin, the dynamics associated with protrusion of
lamellipodia, for example, will now be more accessible
to experimentation, since polymerisation and retrograde flow can be monitored directly. Likewise, the
fluorescently labelled living cell provides a standard
against which methods of fixation and extraction for
correlative biochemical and structural studies may be
more effectively controlled. Fluorescent derivatives of
actin-associated proteins as well as of proteins resident
at actin-membrane interfaces will facilitate investigations of their functions, in relation to actin cytoskeleton
assembly and the rearrangements induced via the
Rho-family signaling pathways.
ACKNOWLEDGMENTS
We thank Dr. Rob Cross for permission to include
data collected on the two-photon and confocal microscope in the Marie Curie Institute and Dr. M. Gimona
for providing purified chicken gizzard vinculin and
discussion. Provision of different probes and cells is also
acknowledged from the following colleagues: Prof. J.
Wehland, Dr. A. Sechi, and Mr. M. Geese (Braunschweig, Germany), Prof. B.A. Imhof, Dr. C. Ballestrem,
and Dr. B. Wehrle-Haller (Geneva, Switzerland), Prof.
H. Faulstich (Heidelberg, Germany), Dr. R. Salgia and
Dr. J. Griffin (Boston, MA, USA). Figure 7 was kindly
provided by Dr. T. Svitkina (Madison, WI, USA). We
thank also Ms. M. Schmittner for photography and Ms.
E. Eppacher for text processing. The instrumentation
used for data collection was funded in part from the
Austrian Science Research Council, the Austrian National Bank and the Seegen Stiftung of the Austrian
Academy of Sciences.
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