CORRECTION
Correction: Actin visualization at a glance
Michael Melak, Matthias Plessner and Robert Grosse
There was an error published in J. Cell Sci. 130, 525-530.
Unfortunately, the diagram of the actin filament in the poster was accidentally reflected during the creation of the figure. As a result, the
filament was illustrated as a left-handed rather than a right-handed F-actin helix in the original version of this article. The online version of
the article has been corrected accordingly. This change has no impact on the concepts presented in this article.
The authors apologise to the readers for any confusion that this error might have caused.
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Journal of Cell Science
CELL SCIENCE AT A GLANCE
Actin visualization at a glance
Michael Melak, Matthias Plessner and Robert Grosse
*
ABSTRACT
Actin functions in a multitude of cellular processes owing to its ability
to polymerize into filaments, which can be further organized into
higher-order structures by an array of actin-binding and regulatory
proteins. Therefore, research on actin and actin-related functions
relies on the visualization of actin structures without interfering with
the cycles of actin polymerization and depolymerization that underlie
cellular actin dynamics. In this Cell Science at a Glance and the
accompanying poster, we briefly evaluate the different techniques
and approaches currently applied to analyze and visualize cellular
actin structures, including in the nuclear compartment. Referring
to the gold standard F-actin mar ker phallo idin to stain actin in
fixed samples and tissues, we highlight methods for vis ualization
of actin in living cells, which mostly apply the principle of
genetically fusing fluorescent proteins to different actin-binding
domains, such as LifeAct, utrophin and F-tractin, as well as anti-
actin-nanobody technol ogy. In addit ion, the compoun d SiR-actin
and the expression of GFP–actin are also applicable for various
types of live-c ell analyses. Overall, the visualiz ation of actin within
a physiological cont ext requires a careful choic e of method, as
well as a tight cont rol of the amount or the expressi on level of a
given detection probe in ord er to minimize its influence on
endogenous actin dynamics .
KEY WORDS: Actin dynamics, Actin probes, Live-cell imaging,
Nuclear actin
Institute of Pharmacology, Biochemical-Pharmacological Center (BPC), University
of Marburg, Karl-von-Frisch-Straße 1, Marburg 35043, Germany.
*Author for correspondence (robert.grosse@staff.uni-marburg.de)
R.G., 0000-0002-3380-5273
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Introduction
Actin is one of the most abundant proteins in eukaryotic cells and its
amino acid sequence is very highly conserved from yeast to man.
Concentrations of actin in euk aryotic cells are well within the
micromolar range (Wu and Pollard, 2005), and, hence, actin fulfills
a multitude of cellular functions, such as intracellular transport,
adhesion and con traction, membrane dynamics and migration,
cytokinesis and cell– cell contact regulation, polarity and cell shape
control (Dominguez and Holmes, 2011), as well as gene regulation
(Louvet and Percipalle, 2008; Bunnell et al., 2011) and other less-
well explored functions in the nucleus (Grosse and Vartiainen,
2013; Plessner and Grosse, 2015).
In contrast to microtubules, which typically form long and
straight cylindrical polymers with an outer diameter of 25 nm
(Wade, 2007), actin filaments (F-actin) are composed of two twisted
helices with a diameter of ∼5 to 9 nm (Holmes et al., 1990). One
intrinsic characteristic of single actin proteins is their highly
dynamic turnover between a monomeric and a polymeric state,
which can result in rapid filament assembly and disassembly.
Although methods for the visualization of single microtubules
have been well established for decades (Sherwin and Gull, 1989;
Kikkawa et al., 1994), it has been and still is a challenge to visualize
specific actin structures and their related functions, in particular
under real-time conditions in living cells and organisms.
Consequently, many different approaches and probes have been
developed over the past years to monitor the dynamics of actin
assembly and actin filament structures in several different model
systems, while mostly trying to illuminate endogenous actin
proteins.
Recently, several publications have discussed in great detail the
benefits and disadvantages of different fluorescent actin reporters
(Belin et al., 2014; Spracklen et al., 2014; Lemieux et al., 2014; Du
et al., 2015). In this short article, and the accompanying poster, we
present an at-a-glance view of the current approaches to visualize
actin filaments, emphasizing the advantages and pitfalls of the
available tools to investigate F-actin not only in the cytoplasm, but
also in the somatic cell nucleus.
Visualization o f F-actin in fixed cells and tissues
Visualization of the actin cytoskeleton was initially performed by
immunofluorescence techniques that used antibody staining against
fixed actin structures (Lazarides and Weber, 1974). To overcome
possible drawbacks of antibody-based labeling techniques, such
as non-specific binding, actin scavenging, high background, or
variations in the epitope among different species, additional actin-
binding components with a high specificity for F-actin were
developed.
The gold standard marker for labeling endogenous actin
filaments in fixed samples still remains phalloidin, a fluorescently
labeled derivative of the phallotoxin, which binds to F-actin with
high affinity (Wulf et al., 1979; Vandekerckhove et al., 1985) (see
poster). Usage of phalloidin is largely restricted to the labeling of
fixed cells, as this toxin only has a low permeability for cell
membranes and more importantly stabilizes actin filaments in vivo
and in vitro (Coluccio and Tilney, 1984; Visegrády et al., 2004;
Wehland et al., 1977). Thus, the actin structures visualized by
fluorescent phalloidin are in most cases arrested or altered to a
certain degree by the necessary fixation and permeabilization
procedures and, therefore, the obtained results need to be interpreted
with caution. Depending on the permeabilization and fixation
protocol, formation of artifacts can occur. For example, it has been
shown that methanol fixation methods are not to be suitable for actin
staining with phalloidin, as it can destroy the native F-actin
conformation, resulting in labeling of artificial structures (Kellogg
et al., 1988). Another drawback of using fluorescent phalloidin to
label endogenous F-actin in native cells and tissues is that it might
not bind to all the actin structures present. For example, the binding
of cofilin to F-actin has been shown to interfere with fluorescent-
dye-labeled phalloidin staining due to alterations of the F-actin
structure (Bunnell et al., 2011; McGough et al., 1997). Finally, at
least seven actin subunits are necessary for successful binding of
phalloidin to F-actin (Kristó et al., 2016), preventing labeling of
short F-actin pol ymers with phalloidin.
Ectopic expression of epitope -tagged actin (i.e. Myc-, HA- or
Flag-tagged actin) in combination with immunofluorescence
staining has been widely used to study actin organization in cells
(Copeland and Treisman, 2002; Miralles et al., 2003). This simple
approach can be helpful, but only if the expression levels of the
probe are carefully controlled and titrated. Clearly, overexpression
of tagged actin variants can be problematic, as even a subtle change
in the amount of global actin can interfere with the physiological
actin dynamics, and could trigger actin polymerization by itself
(Mounier et al., 1997; Ballestrem et al., 1998). In addition, this
approach is not well suited for live-cell imaging owing to the
necessary immunofluorescence staining, unless actin is labeled with
directly stainable tags, such as SNAP- (Keppler et al., 2004;
Lukinavi
cius et al., 2013) or tetracysteine tags (Griffin et al., 1998).
For instance, actin fused to a tetracysteine tag can be directly
visualized by using different labeling reagents, such as the
biarsenical dyes FlAsH and ReAsH (Martin et al., 2005). The
relatively small size of the tetracysteine tag (∼2 kDa) might not
interfere dramatically with different functions of actin, although it
has been shown that the fission yeast formin Cdc12p, which is
required for formation of the contractile ring, excludes tetracysteine-
tagged fission yeast actin, suggesting very stringent structural
requirements for formins to promote the elongation of actin
filaments (Chen et al., 2012). Moreover, owing to potential
chemical toxicity caused by the labeling reaction, such as the
accumulation of FlAsH in active mitochondria (Langhorst et al.,
2006), this technique might be better suited for short-term live-cell
imaging.
Live-cell imaging using GFP–actin derivatives
Tagging actin with GFP derivatives is extensively used for live-cell
analysis. In particular, GFP–actin is suitable for fluorescence
recovery after photobleaching (FRAP) approaches to study actin
dynamics and turnover within a given cellular F-actin structure,
including, for instance, the actin cortex ( Clark et al., 2013) or
protrusions such as lamellipodia (Koestler et al., 2009) (see poster).
However, the relatively large size (∼28 kDa) of GFP-like proteins
(Sliogeryte et al., 2016) can give rise to problems in terms of
incorporation of GFP-tagged actin monomers into filaments. Thus,
the GFP–actin-derived localization pattern might only reflect a part
of the total F-actin network in cells. For example, it has been shown
in yeast that GFP–actin does not incorporate effectively in a specific
subpopulation of actin filaments (Doyle and Botstein, 1996).
Furthermore, certain actin nucleators, such as members of the
formin family, might exclude GFP–actin monomers from their
nucleation and elongation mechanisms, probably due to steric
hindrances during the formation of actin seeds or during efficient
barbed-end actin incorporation (Wu and Pollard, 2005; Vavylonis
et al., 2006; Carvalho et al., 2009).
In summary, as is the case for any genetically encoded protein
tagging, the expression of GFP–actin needs to be accurately
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controlled in any given cellular system. Nevertheless, GFP–actin
has been proven to be highly useful to image actin functions in
living cells or organisms, such as in mammalian cells (Ballestrem
et al., 1998), yeast (Doyle and Botstein, 1996), Dictyostelium
discoideum (Neujahr et al., 1997; Westphal et al., 1997),
Drosophila (Verkhusha et al., 1999) and mice (Gurniak and
Witke, 2007).
Imaging F-actin structures and dynamics using actin-binding
domains
There are currently several probes available to visualize endogenous
actin dynamics in living cells. With the exception of SiR-actin (see
below), these are based on genetically encoded actin-binding
domains (ABDs) that have been derived from yeast or human
proteins, which are fused to GFP derivatives.
The most popular and widely used such tool is LifeAct, a short
17-amino-acid peptide from yeast Abp140 ( Riedl et al., 2008).
There are numerous publications describing the successful
application of LifeAct, in particular in mammalian cell motility
(Rullo et al., 2012; Grikscheit et al., 2015; Suarez et al., 2015; Duleh
and Welch, 2012; Fiolka et al., 2012) (see poster). However,
experimental and technical drawbacks in using LifeAct as an actin
probe in living cells have also been described. For exampl e, one
disadvantage of LifeAct–GFP variants is thei r relatively high
background fluorescence owing to their high affinity for globular
actin (G-actin) (Riedl et al., 2008).
Furthermore, although LifeAct has been suggested to be a
universal marker for actin (Riedl et al., 2008), it is unable to stain
certain actin structures, including a specialized class of actin-based
filopodia in mesenchymal cells of the developing limb bud of chick
embryos (Sanders et al., 2013) and stress-induced cofilin-bound
F-actin structures in mouse striatal neuron-derived STHdh cells
(Munsie et al., 2009). Both studies suggest that extensive coating of
actin filaments with actin-binding proteins, such as cofilin, result in
conformational changes of F-actin, thereby preventing LifeAct from
accessing its binding site. In addition, at higher expression levels,
LifeAct can influence the formation of actin filaments. It has been
reported that, in contrast to expression in somatic cells or weak
expression in the germline, a strong expression of LifeAct in the
Drosophila germline results in severe actin remodeling and fertility
defects (Spracklen et al., 2014). Moreover, it has been shown that
LifeAct affects actin assembly during endocytosis and cytokinesis
in fission yeast (Courtemanche et al., 2016). Although the
underlying mechanisms are not well understood, both studies
claim that the LifeAct-me diated effects on actin dynamics are
concentration-dependent and thus suggest that the use of LifeAct
requires a more elaborate optimization process. Nevertheless, an
important advantage of LifeAct is its relatively low toxicity when its
expression level has been carefully optimized. This is reflected in
the successful generation of transgenic animals, such as mice or
zebrafish (Riedl et al., 2010; Mizoguchi et al., 2016; Phng et al.,
2013; Schachtner et al., 2012). Furthermore, LifeAct has been
successfully used for actin imaging in plants such as Arabidopsis
(van der Honing et al., 2011; Vidali et al., 2009) and in several fungi
(Berepiki et al., 2010; Delgado-Álvarez et al., 2010).
In addition to LifeAct, the tandem calponin homology domains
(CH1 and CH2) of utrophin (UtrCH) are frequently used as a marker
for stable and dynamic F-actin, both in living and fixed cells (Burkel
et al., 2007). UtrCH comprises the first 261 amino acids of human
utrophin, an actin-binding, dystrophin-related protein (Winder et al.,
1995). It has been suggested that the two CH domains of Utr261
interact with adjacent actin subunits (Lin et al., 2011). As actin is
highly conserved, UtrCH has been successfully used as a marker for
actin filaments in a wide range of organisms and cell types,
including mammali an, amphibic or echinodermal oocytes and
embryos, without showing negative effects of UtrCH on both actin
organization and dynamics (Burkel et al., 2007; Holubcová et al.,
2013) (see poster). However, more severe defects related to actin
dynamics have been reported, such as cortical actin breakdown and
alterations in the morphology of actin bundles, as well as female
sterility during Drosophila oogenesis, when UtrCH is expressed at
higher levels (Spracklen et al., 2014).
Another tool, later on termed F-tractin, is a 43-amino-acid long
peptide from the rat actin-binding inositol 1,4,5-trisphosphate 3-
kinase A [corresponding to residues 10–52 (Belin et al., 2014)]
(Schell et al., 2001) (see poster). F-tractin is less-well studied than
LifeAct and thus there are fewer imaging studies using this probe.
Because F-tractin is larger in size than LifeAct, it might interfere
with other regulatory or accessory F-actin-binding proteins. For
instance, F-tractin expression has been shown to alter the
organization and morphology of the actin cytoskeleton in a
mesoderm-derived cell line from Xenopus (Belin et al., 2014).
However, in contrast to LifeAct and UtrCH, F-tractin has been
reported to not perturb actin rearrangement during Drosophila
follicle development (Spracklen et al., 2014), suggesting that it
might be the most suitable actin probe to study F-actin dynamics
in Drosophila nurse cells. However, further studies are necessary to
fully evaluate the potential advantages of using F-tractin over the
currently more frequently used imaging probes.
Furthermore, as mentioned above, some actin markers are
excluded from specific actin filament structures, and therefore
distinct actin probes might display a differential cytoskeletal
architecture. An interesting recent study compared the distribution
of GFP–actin, F-tractin, LifeAct and utrophin in different living
cells (Belin et al., 2014) and found, for example, that actin-binding
probes often insufficiently stain actin in filopodia-rich regions,
whereas GFP– actin is incorporated well into filopodia. By contrast,
GFP–actin is often excluded from stress fibers and lamellar filament
structures, reflecting its incompatibility with other endogenous
actin-binding proteins. Overall, the authors concluded that among
all the analyzed fluorescent derivatives of actin and ABDs, F-tractin
most accurately reproduced F-actin structures as they could be
visualized by phalloidin labeling in a wide variety of cells (Belin
et al., 2014).
More recently, the cell-permeable chemically synthesized probe
SiR-actin, has been introduced (D’Este et al., 2015); it is structurally
derived from the high affinity F-actin-binding toxin Jasplakinolide
from the marine sponge Jaspis johnstoni (Bubb et al., 1994) (see
poster). Advantages of SiR-actin are its ease of use in imaging
approaches as it circumvents any time-consuming cell transfection
and protein overexpression systems. However, caution must be
taken in live-cell imaging as SiR-actin might cause F-actin
stabilization or induce actin polymerization owing to its structural
similarities to Jasplakinolide. This can make interpretations of
observed F-actin structures more difficult, as they might have been
assembled by the probe itself. Further studies are therefore needed to
fully assess the advantages and possible limitations of SiR-actin
over more established actin probes.
Visualizing actin dynamics using actin-directed nanobodies
A more novel approach to visualize actin filame nt assembly and
disassembly in living cells utilizes single-domain antibodies, or
nanobodies, which specifically bind to a singular antigen; these
nanobodies are composed of a single domain and do not rely on a
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quaternary structure as is the case for conventional antibodies. One
example is the Actin-Chromobody (ChromoTek), a cameloi d
nanobody directed against actin that has been genetically encoded
and fused to TagGFP2 (see poster). When expressed in cells, the
Actin-Chromobody also produces a high level of background
fluorescence, similar to LifeAct. However, in contrast to LifeAct,
the Actin-Chromobody does not seem to influence apparent actin
dynamics, even when it is expressed at high levels. So far, the Actin-
Chromobody has been successfully applied to visualize actin in
mammalian nuclei (Plessner et al., 2015), in tobacco leaf cells
(Rocchetti et al., 2014) and zebrafish (Panza et al., 2015).
Nevertheless, at present, it remains a novel and less-well explored
tool.
Nuclear probes for monitoring F-actin assembly
Nuclear actin filaments and their specific functions are an emerging
topic in the field of actin research. However, there are several
reasons why the visualization of actin filaments in the nucleus is
particularly difficult. First, the concentration of actin in the nucleus
is much lower than in the cytoplasm (Baarlink et al., 2013). Second,
the assembly of nuclear actin filaments is only triggered by specific
signaling cues, for example in response to extracellular signals, such
as serum factors (Baarlink et al., 2013) or DNA damage (Belin et al.,
2015), or in the context of integrin-based cellular adhesion and
mechanotransduction (Plessner et al., 2015). Finally, actin, as well
as most of the actin-regulatory proteins found in mammalian cell
nuclei, dynamically shuttles between the cytoplasm and the nuclea r
compartment (Kristó et al., 2016).
Staining of fixed nuclear actin filaments can be achieved by
careful fixation with formaldehyde or glutaraldehyde, followed by
phalloidin staining (Baarlink et al., 2013) and image acquisition
with a state-of-the-art confocal microscope as well as a sensitive
detection method (see poster). Live-cell imaging of nuc lear actin
requires the expression of actin-detecting probes that are fused to a
nuclear localization signal (NLS) (see poster), such as the nuclear-
targeted Actin-Ch romobody–GFP (nAC–GFP) (Plessner et al.,
2015). This helps to circumvent the comparably high signal of actin
that emanates from the cytoplasm, and, indeed can be used to
visualize nuclear actin filament assembly in living cells (Plessner
et al., 2015). However, it is important to note that a compartment-
specific localization of actin-detecting probes might influence the
shuttling dynamics of actin and actin-binding proteins. Thus, we
recently generated modified nuclear actin probes that also contains
an additional nuclear export signal (NES) (LifeAct–GFP–NLS–
NES and Actin-Chromobody–TagGFP2–NLS–NES), which allow
for dynamic nucleocytoplasmic shuttling of the probe itself and so
additionally enable the simultaneous visualization of cytoplasmic
and nuclear actin structures (Plessner et al., 2015).
It is, therefore, important to avoid artificial transient
overexpression to ensure that the probe does not interfere with
nuclear actin dynamics (Du et al., 2015). In general, usage of stably
expressing cell lines or viral expression systems might be better suited
to avoiding artifacts caused by genetically encoded actin probes.
Conclusions an d perspectives
A number of approaches and probes have been generated to
visualize actin and the many types of filamentous structures it
generates. The approaches and tools discussed here each have their
advantages and pitfalls, and some of the too ls are currently more
commonly employed than others. One limitation of genetically
encoded actin probes as well as phalloidin is their inability to
distinguish between specific actin isoforms. Antibody-based
approaches have been developed to address this (Perrin and
Ervasti, 2010).
A major reason to decide for or against a particular actin-detection
probe is the cellular context under study. Using the example of
LifeAct, it has been proposed to be the most suitable actin probe to
visualize the dynamic rearrangement of the actin cytoskeleton in
Dictyostelium, as it labels a more complete subset of actin structures
than other actin-binding probes (Lemieux et al., 2014). The
application of LifeAct was also favored over actin fused to
fluorophore in the context of visualizing actin rearrangements
during cellular mechanotransduction events (Sliogeryte et al.,
2016; Deibler et al., 2011). In contrast, LifeAct should be used
with caution for actin visualization in fission yeast, as it can cause
severe concentration-dependent defects in endocytosis and
cytokinesis (Courtemanche et al., 2016). Furthermore, the
suitability of the actin probe UtrCH is also highly dependent on the
model organism or cell line. For example, UtrCH has been
successfully used to visualize actin dynamics in mouse (Schuh,
2011; Holubcová et al., 2013) and Xenopus oocytes, as well as in
several species of Echinodermata (Burkel et al., 2007). In contrast, a
strong expression of UtrCH in the Drosophila germline has been
reported to disrupt reorganization of the actin cytoskeleton during
Drosophila oogenesis (Spracklen et al., 2014).
In addition, the choice of an appropriate fluorescent protein to be
fused to an ABD is also of importance. For instance, a recent
publication has demonstrated that the subset of F-actin that is
recognized by diverse ABDs can be altered by the fluorescent
protein or the linker sequence between the ABD and the fluorescent
protein (Lemieux et al., 2014). New applications for fluorescent
labeling of proteins have become available recently, such as the use
of Y-FAST, a small monomeric protein tag enabling reversible
binding and activation of a cell-permeant and nontoxic fluorogenic
molecule (Plamont et al., 2016), or the flavoprotein improved LOV
(iLOV), a fluorescent protein based on the light, oxygen or voltage
domain (LOV) from a variety of sources (Buckley et al., 2015). Both
molecules offer advantages over GFP and other related fluorescent
reporters owing to their relatively small size. Therefore, they might
provide valuable alternatives for actin visualization when fused to
actin itself or to ABDs.
In summary, every detection method has limitations and might
alter endogenous actin dynamics to various degrees. Hence, the
choice and usage of any given probe has to be carefully considered
and evaluated in the cellular system under study to be able to achieve
an optimal reflection of the actin structure of interest within its
physiological context. It is therefore important to validate obtained
results with several probes to minimize any potential non-
physiological observations or formation of artificial F-actin
aggregations caused by the probe itself.
Acknowledgements
We thank members of our laboratory for helpful discussions. We thank J. Ivaska for
providing mApple-F-tractin.
Competing interests
The authors declare no competing or financial interests.
Funding
This work was supported by grants from the Deutsche Forschungsgemeinschaft
(GR2111-7-1) and the Wilhelm Sander-Stiftung (2013.149.1) to R.G.
Cell science at a glance
A high-resolution version of the poster and individual poster panels are available for
downloading at http://jcs.biologists.org/lookup/doi/10.1242/jcs.189068.
supplemental
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