ß 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
COMMENTARY
ARTICLE SERIES: IMAGING
Molecular probes to visualize the location, organization and
dynamics of lipids
ABSTRACT
Cellular lipids play crucial roles in the cell, including in energy
storage, the formation of cellular membranes, and in signaling and
vesicular trafficking. To understand the functions and characteristics
of lipids within cells, various methods to image lipids have been
established. In this Commentary, we discuss the four main types
of molecular probes that have significantly contributed to our
understanding of the cell biology of lipids. In particular, genetically
encoded biosensors and antibodies will be discussed, and how they
have been used extensively with traditional light and electron
microscopy to determine the subcellular localization of lipids and
their spatial and temporal regulation. We highlight some of the
recent studies that have investigated the distribution of lipids
and their ability to cluster using super-resolution and electron
microscopy. We also examine methods for analyzing the
movement and dynamics of lipids, including single-particle
tracking (SPT), fluorescence recovery after photobleaching
(FRAP) and fluorescence correlation spectroscopy (FCS).
Although the combination of these lipid probes and the various
microscopic techniques is very powerful, we also point out several
potential caveats and limitations. Finally, we discuss the need for
new probes for a variety of phospholipids and cholesterol.
KEY WORDS: Electron microscopy, Imaging, Lipid, Superresolution
Introduction
With more than1000 lipid species in the typical mammalian cell,
understanding their functions is a daunting task and requires an
armamentarium of techniques and probes. Although some classes
and species of lipids serve major structural roles within the cell,
others are potent signaling molecules kept at low concentrations.
With so many individual molecules and species of lipids within
the cell, the level of organization is truly remarkable. Not only
does lipid composition vary between the specific organelles
within the cell, but there also exists an asymmetric distribution
between leaflets of the same bilayer and inhomogeneous
distribution within the same leaflet. This organization is the
result of a multitude of lipid–lipid and lipid–protein interactions,
1
Keenan Research Centre for Biomedical Science, St. Michael’s Hospital, 209
Victoria Street, Toronto, ON M5S 1T8, Canada. 2Department of Surgery,
University of Toronto, Toronto, ON M5T 1P5, Canada. 3Department of
Biochemistry, University of Toronto, Toronto, ON M5S 1A8, Canada. 4Institute of
Medical Science, Faculty of Medicine, University of Toronto, Toronto, ON M5S
1A8, Canada. 5Institute for Biomedical Engineering and Science Technology
(iBEST) at Ryerson University and St. Michael’s Hospital, Toronto, ON M5B 2K3,
Canada.
*Author for correspondence ([email protected])
metabolic pathways, lipid transporters and vesicular transport
pathways.
Lipids perform three general functions within eukaryotic cells:
they (1) operate as energy stores in the form of triacylglycerides
and cholesterol esters, (2) act as building blocks of cellular
membranes owing to their amphiphilic nature, and (3) serve as
secondary messenger and/or regulators in signal transduction and
transport processes (van Meer et al., 2008). Owing to the array of
functions and their biological importance, a great deal of time and
effort has gone into understanding, not only these lipids, but also
the enzymes and proteins that regulate their metabolism and
subcellular distribution and function. Collectively, this research
constitutes the field of lipidomics. Lipidomics has been greatly
bolstered since the early 2000s owing to advances and availability
in mass spectrometry, computational methods and microscopy.
The ability to quantify lipids using mass spectrometry and
to determine their localization in a dynamic fashion represent
powerful complementary techniques.
Mass spectrometry enables us to not only perform accurate
quantitative analysis of lipid classes but also allows us to
distinguish molecular species of lipids due to variations in their
acyl chains (Han and Gross, 2003; Watson, 2006). When
combined with subcellular fractionation, mass spectrometry can
catalog the lipidome of specific organelles. This approach,
although very powerful, does have a variety of limitations. For
instance, in many of these experiments a large number of cells are
required for robust data generation, and it takes several hours
to isolate the organelles or membranes of interest. This has led
researchers to develop techniques in which spatial information is
added to the mass spectrometry data. Advances in this area have
led to mass-spectrometry-based imaging approaches. One
such technique uses matrix-assisted laser desorption ionization
(MALDI) mass spectrometry together with controlled twodimensional movement of the sample during the data collection
(Cornett et al., 2007). Thus, it enables the results for specific lipid
species to be represented as an image. The major drawback of this
technique is that, owing to limited resolution, it is more suitable
for tissue slices than individual cells. Alternative approaches
have been developed that achieve sub-micron resolution and
therefore are more desirable for cell biologists. Recent technical
developments in time-of-flight, secondary ion mass spectroscopy
(SIMS) have enabled the analysis of both the composition and
organization of lipids with a lateral resolution of ,500 nm
(Touboul et al., 2011). The use of next-generation SIMS and
incorporating the use of stable-isotope-labeled lipids has allowed
researchers to obtain a lateral resolution of 100 nm for a few
classes of lipids (for more information, see Kraft and
Klitzing, 2014). Although these are great tools, it remains
difficult to examine dynamic and asynchronous processes,
such as cell migration or macropinocytosis in cell populations.
This is especially true for low-abundance lipids, such as
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Masashi Maekawa1 and Gregory D. Fairn1,2,3,4,5,*
COMMENTARY
Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
phosphoinositides and diacylglycerol (DAG), that can act as
potent signaling molecules but have short lifetimes and precise
sites of action. Furthermore, mass spectrometry has limited
ability to elucidate the transbilayer distribution of phospholipids
and cholesterol in the inner (or cytosolic) and the outer (or
luminal) leaflets of cellular membranes or other membrane areas
with spatial inhomogeneities.
Microscopic visualization of lipids represents a powerful
technique that is complementary to mass-spectrometry-based
analysis. However, with the exception of a few lipid molecules,
such as dehydroergosterol and cholestatrienol, the vast majority
of lipids are not intrinsically fluorescent (Rogers et al., 1979).
Therefore a variety of surrogates and probes have been identified
or generated to facilitate the microscopic analysis of lipids. These
probes can be combined with other lipid probes of varying
spectral properties or with fluorescent proteins for colocalization
studies. In addition, many of these probes are suitable for live-cell
imaging and are very useful for monitoring the subcellular
localization and dynamics of specific lipids. Notably, this method
enables the monitoring of specific lipids in processes such as
phagocytosis and cell polarization (Yeung et al., 2008; Fairn
et al., 2011a; Sarantis and Grinstein, 2012). However, one
limitation of this approach is that, at the current time, these
probes cannot distinguish the acyl chains of glycerolipids. Thus to
provide unambiguous conclusions, we recommend performing
complementary studies (i.e. using both mass-spectrometry and
visualization of lipids) whenever possible.
This Commentary will review approaches for imaging lipids
using specific lipid probes, including a discussion of the types of
available probes and their use with a variety of conventional and
super-resolution imaging techniques. First, we discuss general
approaches for analysis of intracellular ‘localization’ of lipids
using fluorescent microscopy, before moving on to advanced
methods for visualization of lipids in clusters with electron
microscopy and sub-diffraction-limited microscopy. Finally,
we highlight the probes and techniques that are being used to
analyze the dynamics of lipids by fluorescence recovery after
photobleaching (FRAP), fluorescence correlation spectrometry
(FCS) and single-particle tracking (SPT).
Lipid probes and their subcellular localization
For imaging of lipids with standard epifluorescence or confocal
microscopy, the selection of suitable probes is very important and
can vary owing to experimental design. A variety of established
lipid probes and surrogates (Table 1) have been used to generate
a map of many of the lipids in the cell (Fig. 1). In general, probes
used for the visualization of lipids can be divided into four
categories: (1) fluorophore-labeled lipids, (2) antibodies, (3) toxin
domains, and (4) genetically encoded protein domains. Each of
them has advantages and disadvantages associated with their use,
and investigators must keep in mind that antibodies and protein
domains must compete with endogenous lipid binding proteins to
localize to the membrane. In this regard, many of these proteins
only detect the ‘free’ or available lipids. That said, these modular
domains have been used extensively to determine the distribution
of lipids between organelles and membrane leaflets.
Fluorophore-conjugated lipids
Fluorophore-conjugated lipids are a conceptually attractive way
to examine the movement and localization of the particular lipids
Lipid
Probes (Kd)
Intracellular localization
References
PtdSer
Anti-PtdSer antibody
Lact-C2 (190 nM)
Annexin Va
TopFluor-PS
Sos1-PH (470 nM)
2PABD
PKCd-C1
Lysenin (5.3 nM)
BODIPY–sphingomyelin
Anti-ceramide antibody
BODIPY–ceramide
BODIPY–lactosylceramide
Anti-LBPA antibody
Filipin
TopFluor–cholesterol
Perfringolysin O-D4
26EEA1-FYVE (38 nMd)
p40phox-PX (0.71 nM)
Osh2-PH (1.3 mM)
FAPP1-PH
PLCd-PH (210 nMe)
TAPP-1-PH (5.4 nM)
GRP-1-PH (27.3 nMe)
Btk1-PH
Akt-PH [570 nM for PtdIns(3,4)P2;
400 nM for PtdIns(3,4,5)P3]
PM; nascent phagosomes
PM (inner); endosomes
PM (outer)
PM; endosomes
PM (inner)b
Membrane rufflesc
Golgi; PM (inner); phagosomes
PM (outer)
PM; endosomes; Golgi
PM; Golgi
Golgi
PM; endosomes; Golgi
Late endosomes
PM; endosomes
Endosomes; Golgi
PM (outer)
Early endosomes
Yeung at al., 2006
Yeung et al., 2008
Andree et al., 1990
Kay et al., 2012
Zhao et al., 2007
Bohdanowicz et al., 2013
Colón-González and Kazanietz, 2006
Yamaji et al., 1998
Puri et al., 2001
Krishnamurthy et al., 2007
Pagano et al., 1991
Puri et al., 1999
Kobayashi et al., 1998
Börnig and Geyer, 1974
Kleusch et al., 2012
Shimada et al., 2002
Gillooly et al., 2000
Ellson et al., 2001; Stahelin et al., 2003
Roy and Levine, 2004; Yu et al., 2004
Godi et al., 2004
Stauffer et al., 1998
Dowler et al., 2000
Gray et al., 1999
Varnai et al., 1999
Frech et al., 1997; Gray et al., 1999
Phosphatidic acid
DAG
Sphingomyelin
Ceramide
Lactosylceramide
LBPA
Cholesterol
PtdIns3P
PtdIns4P
PtdIns(4,5)P2
PtdIns(3,4)P2
PtdIns(3,4,5)P3
PtdIns(3,4)P2 and PtdIns(3,4,5)P3
a
PM (inner)
Golgi
PM (inner)
Membrane rufflesc
Membrane rufflesc
Membrane rufflesc
Ca2+ is required;
during serum stimulation;
c
during phagocytosis and macropinocytosis;
d
Kd of GST–FYVE;
e
measured using free inositol phosphates (Lemmon et al., 1995; Kavran et al., 1998). LBPA, lysobisphosphatidic acid.
b
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Table 1. Established lipid probes
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Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
A
C
B
D
Medium or lumen
Cytosol
Fluorophore-labeled lipids
Domain of toxins
Domain of genetically
encoded protein
Fluorophore-labeled
second antibody
Fluorophore
Fluorophore
Anti-LBPA
Bodipy–LacCer
E
Antibody for lipids
His–GFP–PFO-D4
(Cholesterol)
Medium
PM
GFP–PLCδ-PH
[PtdIns(4,5)P2]
Membrane ruffles
SM, Cholesterol
PtdSer
PA
PtdIns(3,4,5)P3
PtdIns(3,4)P2
PtdSer, PtdIns(4,5)P2
PtdSer
PtdIns3P
cis-Golgi
Early endosomes
PtdSer
LBPA
PtdIns(3,5)P2
DAG
PtdIns4P
LacCer
Late endosomes
PtdSer
Recycling endosomes
Fig. 1. Subcellular distribution of lipids visualized by four types of lipid probes. (A–D) The schematics at the top illustrate the different types of commonly
used lipid probes and include an example below. Note that the size of lipids and probes (especially, antibodies and toxin domains) are not to scale. Additionally,
antibodies and some toxins have multivalent binding sites which enable them to bind to more than one lipid at a time. (A) BODIPY–lactosylceramide (LacCer) is
used as a probe for lactosylceramide, which localizes to the trans-Golgi network (TGN). Representative image of a cell after BODIPY–LacCer was added
to the medium, followed by a 60-minute chase period in label-free medium. The yellow dotted line outlines the cell. (B) LBPA (green in the schematic) can be
visualized with anti-LBPA antibodies. Here, fixed CHO cells were incubated with mouse monoclonal anti-LBPA antibody (6C4) in the presence of 0.05% saponin,
followed by incubation with fluorophore-labeled secondary antibody. As shown in the image, it localizes to late endosomes. The yellow dotted line indicates the
outline of the cell. (C) Cholesterol (orange in the schematic) localizes in the outer leaflets of PM as visualized by using the cholesterol-binding toxin Perfringolysin
O (PFO). For the image, domain 4 of the PFO protein was purified (66His-tagged PFO-D4), added to cells and imaged using confocal microscopy.
(D) Phosphoinositides (blue in the schematic) can be visualized by using genetically encoded protein domains. In the example shown here, PtdIns(4,5)P2 in the
inner leaflets of PM was visualized by GFP–PLCd-PH in CHO cells using confocal microcopy. Scale bars: 10 mm. (E) Schematic diagram of the intracellular
distribution of lipids in the cytosolic (inner) leaflets and the luminal (outer) leaflets of cellular membranes. The subcellular distribution of lipid in different cellular
membranes varies throughout the cells. Presumably, the luminal leaflets of intracellular compartments illustrated contain a certain degree of cholesterol and
sphingomyelin (SM, highlighted in orange). PA, phosphatidic acid.
within the cell. They are easily delivered to cells by pre-loading
them onto carrier molecules, such as bovine serum albumin (BSA),
and can be added to the growth medium. In general, the
fluorophores are conjugated to either the polar head group (i.e.
Rhodamine-labeled phosphatidylethanolamine) or to one of the
acyl chains or hydrophobic components (i.e. BODIPY–cholesterol)
(Axelrod et al., 1976; Li et al., 2006). One concern is that once the
head group of the phospholipid is modified with a fluorescent
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trans-Golgi
Cytosol
molecule, the resulting phospholipid no longer retains the
properties of the parent molecule. This is also a concern when
the hydrophobic acyl chains are modified with molecules, such as
the low-polarity boron-dipyrromethene (BODIPY) or the more
moderately polar nitro-benzoxadiazolyl (NBD). As a result, the
fluorophore-conjugated lipids likely have different biophysical
properties than the natural lipids and this could impact on their
characteristics, such as the rate of intramembrane transbilayer
movement (so called flip-flop), spontaneous intervesicle or
organelle transfer, and lateral packing (Chattopadhyay and
London, 1987; Martin and Pagano, 1987; Abrams and London,
1993; Martin and Pagano, 1994; Kaiser and London, 1998;
Baumgart et al., 2007; Elvington and Nichols, 2007; Kay et al.,
2012; Sezgin et al., 2012). Although these concerns might not be
crucial for some experiments, it should also be noted that not all of
these molecules are metabolically inert, and it is impossible to
distinguish between the original fluorophore-labeled lipids and
their metabolites that are generated by various enzymes (e.g.
phospholipases) in vivo. Thus, although fluorescently labeled lipids
are still useful for in vitro assays, their in vivo use is somewhat
more limited.
Antibodies
Antibodies have been widely used for determining the
localization and abundance of proteins in immunocytochemistry
and immunohistology. Similarly, there are a few antibodies
that recognize endogenous unlabeled lipids, with a classic
example being the anti-lysobisphosphatidic acid (LBPA)
antibody (Kobayashi et al., 1998). However, the visualization
of lipids with antibodies often requires the cells to be fixed and
permeabilized for the antibody to gain access to intracellular
compartments. Care must be taken, because the fixation and
permeabilization procedure might not only result in unintended
removal and redistribution of lipids but also in differences
in accessibility between different organelles. This concept is
illustrated in a study by Irvine and co-workers that used
formaldehyde and glutaraldehyde to fix and saponin to
permeabilize cells for immunofluorescence detection of
phosphoinositides in the plasma membrane (Hammond et al.,
2009a). Alternatively, to examine phosphoinositides in the Golgi,
the authors used a lower concentration of formaldehyde,
no glutaraldehyde and permeabilized cells using digitonin
(Hammond et al., 2009a). Yet despite this attention to detail, it
has been demonstrated that integral membrane proteins and lipids
are very difficult, if not impossible, to truly fix (Tanaka et al.,
2010). Beyond the issues with fixation, it must be noted that
antibodies are significantly larger than phospholipids and
typically recognize two or more lipid molecules. Thus, it is
unclear whether antibodies only recognize a particular pool of
lipid or whether they are able to recognize all of the lipid present.
One final limitation is that, in general lipids, are poor antigens
owing to their presence in the immunized animal (Alving and
Richards, 1977; Schuster et al., 1979), and currently only a few
antibodies are suitable for the identification of specific lipids,
such as ceramide and LBPA. However, until alternative probes
for these lipids become available, these antibodies constitute a
viable approach for their imaging.
Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
cholesterol-dependent cytolysins (Mizuno et al., 2011). Many of
these toxins – or for safety and cell viability reasons, non-toxic
domains thereof – can be generated in recombinant form,
either as fusion proteins with fluorescent proteins, such as GFP,
or chemically conjugated to molecules such as Alexa Fluor
succinimidyl ester. These probes have been used extensively to
study the outer leaflet of the plasma membrane, as well as
the lumen of endocytic pathway vesicles (Yachi et al., 2012).
Typically, these types of probes are very specific and can be used
for live-cell imaging. However, as with antibodies, their main
limitation is that only a few such toxins have been identified, and,
moreover, they typically recognize lipids that reside within
exofacial leaflet of the PM.
Domains of genetically encoded proteins
Analogous to toxin domains is the use of modular protein
domains as probes for lipids, and several examples are included in
Table 1. These modular domains are genetically encoded proteins
that are often expressed as fusions with GFP or mCherry. This
type of probe has proven very useful for the analysis of not only
the intracellular localization of lipids but also the dynamic
changes in these lipids in response to stimuli (Várnai and Balla,
2006; Schlam et al., 2013; Steinberg et al., 2014). These lipidbinding domains are produced in the cytosol and translocate to
membranes where the ligands are located. Thus quantification of
membrane lipid changes should be measured as the ratio of
fluorescence intensity in the membrane of interest and the
cytosol. Several such probes (i.e. PLCd-PH, Akt-PH and EEA1FYVE) have been used to visualize the distribution and
metabolism of various phosphoinositides in vivo (Várnai and
Balla, 2006). In many of these experiments, binding-deficient
mutant versions of the probe can be used as a negative control.
Alternatively, the levels of the lipid in question can be depleted
by targeted knockdown of the biosynthetic enzyme or by
treatment with pharmacological inhibitors to ensure the probe is
responding properly to the changes. However, caution must be
used when interpreting the results obtained as their affinity and
specificity can vary greatly. Furthermore, highly expressed
probes can potentially bind to a significant fraction of the lipid,
thereby preventing its normal function (Gillooly et al., 2000;
Fairn et al., 2007; Lemmon, 2008).
Collectively, these four types of lipid probes have allowed lipid
researchers and other cell biologists to examine the subcellular
localization of specific lipid classes and determine how lipids and
electrostatics contribute to organelle identity. Beyond the
subcellular distribution of these probes using standard light
microscopy, some of these have been used for high-resolution
electron and super-resolution microscopy. With the increasing
availability of super-resolution instruments and the ever-growing
list of lipid probes, it should be possible to obtain considerably
more detailed information regarding lipid clustering and
their inhomogeneity in the future. In the remainder of this
Commentary, we will highlight a number of important findings
that have been achieved using such probes in combination with
high-resolution and dynamic live-cell imaging techniques to
determine the lateral movement and confinement of lipids.
Electron microscopic imaging of lipids
Toxin domains
Many secreted toxins and pore-forming molecules recognize
lipids in the exofacial leaflet of the plasma membrane (PM),
including the cholera toxin B subunit, lysenin and various
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Electron microscopy (EM) has been used extensively in the field
of cell biology and was a breakthrough technique used in many
pioneering studies. Although EM is not compatible with live-cell
imaging, its high resolution (,0.2 nm) enables the observation of
Journal of Cell Science
COMMENTARY
intracellular structures and localization of molecules in great
detail (Palade, 1952; Palade and Porter, 1954; Knott and Genoud,
2013). However, to date, EM has not been extensively used to
examine the localization of lipids. The major limitation is that
many of the chemical fixation procedures and processing
protocols used to perform EM of proteins do not allow to
sufficient preservation of membrane lipids. However, more recent
protocols that involve the use of rapid freezing followed by
dehydration (freeze substitution) or freeze fracturing make it
possible to maintain phospholipids in their native (or near-native)
state (van Genderen et al., 1991; Voorhout et al., 1991; Mobius
et al., 2002) and they have been used to visualize antibodies,
toxins and genetically encoded probes of lipids in order to obtain
a high-resolution map of the lipids under investigation in cellular
membranes.
In general, two types of methods can be used to monitor lipids
using EM, pre-fixation or post-fixation labeling of lipids.
Although the addition of the probes that recognize the specific
lipids occur at different times, in both cases, the probes are
detected after fixation by using an antibody or protein A
conjugated to colloidal gold. Pre-fixation detection of lipids is
accomplished by using a plasmid-based biosensor that is
expressed in the cytosol as an epitope-tagged fusion to allow
for visualization. Examples of this strategy include the use of a
tandem FYVE domain to visualize phosphatidylinositol 3phosphate (PtdIns3P), the pleckstrin homology (PH) domain
from phospholipase C for phosphatidylinositol 4,5-bisphosphate
[PtdIns(4,5)P2] and the Lact-C2 domain for phosphatidylserine
(PtdSer) (Fairn et al., 2011b; Zhou et al., 2014). During the
chemical fixation process, the protein-based probes are
crosslinked to proteins in their vicinity, and thus maintain their
original distribution. Therefore, even if the lipids of interest are
not maintained during the rapid fixing process, these protein
probes will provide an accurate representation of the lipids. For
example, using the Lact-C2 and AKT-PH to visualize PtdSer
and phosphatidylinositol 3,4,5-trisphosphate [PtdIns(3,4,5)P3],
respectively, EM analysis has demonstrated that both lipids are
enriched in the plasma membrane over other organelles (Fairn
et al., 2011b; Zhou et al., 2014). However, the enhanced
resolution of EM revealed that both of these probes form
clusters in the PM, and that PtdSer is enriched in caveolae (Fairn
et al., 2011b; Zhou et al., 2014). Taken together, these studies
have provided details regarding the localization of lipids that
cannot be obtained with conventional light microscopy. However,
one limitation of this approach is that because the probes are
expressed in the cytosol they only have access to the cytosolic
leaflet of organelles.
Post-fixation labeling and detection offers the advantage that
the probe has access to both leaflets of the membrane bilayer. In
general, the procedures involve rapid freezing, either freeze
substitution and embedding in a resin, or freeze-fracture replica
formation. Following either of these preparations, the cell
sections are labeled with the probe of interest. During this
processing, care must be taken to maintain the lipids in their
proper location while preserving their accessibility to the
recombinant probe (van Genderen et al., 1991; Voorhout et al.,
1991; Mobius et al., 2002; Fujita et al., 2009; Tanaka et al., 2010;
Fairn et al., 2011b). Another advantage of this approach is that
other probes, such as antibodies and potentially toxins, can also
be used to detect the lipid under investigation. For instance, use of
an anti-LBPA antibody has demonstrated that there is an
enrichment of LBPA in late endosomes over early endosomes
Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
(Kobayashi et al., 1998). This approach has also been used to
examine the localization of PtdIns3P and PtdSer using purified
recombinant GST–26EEA1-FYVE and GST–Lact-C2 probes,
respectively (Gillooly et al., 2000; Fairn et al., 2011b). In the
PtdIns3P study, it was shown that a large portion of the PtdIns3P
formed on the limiting membrane of endosomes is internalized
into intraluminal vesicles during multi-vesicular body formation
(Gillooly et al., 2000; Fig. 2A). Owing to the small size of this
endocytic structure, fluorescence microscopy is unable to
distinguish a signal emanating from the limiting membrane or
the lumen of this compartment. Thus, post-fixation labeling of the
PtdIns3P suggests that a large proportion of PtdIns3P could be
consumed during intraluminal body formation (Gillooly et al.,
2000).
Recently, a freeze-fracture labeling approach was used to
examine PtdIns3P distribution during autophagosome formation
in yeast and mammalian cells (Cheng et al., 2014; Fujimoto et al.,
2014). Using the phox homology (PX) domain of p40phox (also
known as NCF4) in yeast, it has been shown that PtdIns3P is
enriched on the luminal leaflet of the autophagosomal isolation
membrane. Surprisingly, when the experiment was performed
with mammalian cells, the PX domain detected PtdIns3P in the
cytosol-facing leaflet (Cheng et al., 2014; Fujimoto et al., 2014).
These observations suggest that although many of the regulators
involved in autophagy are conserved between yeast and humans
the precise molecular mechanisms are possibly different.
Super-resolution microscopy of lipids
Recent developments in fluorescence microscopy techniques
have allowed researchers to overcome the limitations that are
imposed by the diffraction boundary of conventional fluorescence
microscopy. The series of recent advancements in superresolution microscopy has resulted in a decrease in the limits of
optical resolution from ,250 nm to ,10 nm (Galbraith and
Galbraith, 2011). For the imaging of lipids at this resolution, a
variety of ensemble or single-molecule techniques have been
used, For instance, by using photoactivated localization
microscopy (PALM), a recent study has shown that there are
clusters of cholesterol and sphingomyelin in the outer leaflets of
PM by labeling with recombinant domains of the toxins Dronpa–
PFO-D4 and Dronpa–lysenin, respectively (Fig. 2B) (Mizuno
et al., 2011). Making use of stimulated emission depletion
(STED), it could be shown that PtdIns(4,5)P2 is highly enriched
within segregated domains in the PM (Fig. 2C) (van den Bogaart
et al., 2011). In this particular study, PtdIns(4,5)P2 was visualized
using both a recombinant citrine-tagged GST–PLCd-PH
(citrine is a YFP analog) protein and anti-PtdIns(4,5)P2
antibody. Furthermore, a similar study used stochastic optical
reconstruction microscopy (STORM) imaging to demonstrate the
presence of PtdIns(4,5)P2 and PtdIns(3,4,5)P3 in the PM by using
anti-phospholipid antibodies that are directly conjugated to Alexa
Fluor 647 (Wang and Richards, 2012). Collectively, the ability to
visualize lipids using EM and super-resolution microscopy
constitutes a powerful tool to help to better understand the
organization of signaling hubs these lipids are part of.
Approaches to visualize lipid dynamics
Cellular membranes, with the exception of lipid droplets,
are typically lipid bilayers formed by the amphiphilicity of
glycerolipids and cholesterol. The weak hydrophobic interaction
between lipids in membranes enables the lateral diffusion of
lipids and proteins in membranes (Singer and Nicolson, 1972;
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A
Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
EM
GST−2xFYVE
B
PALM
Dronpa−Lysenin
C
Confocal
STED
Anti−PtdIns(4,5)P2 antibody
Holthuis and Levine, 2005; van Meer et al., 2008). As mentioned
above, some lipids form clusters in the PM, and these clusters are
likely to be important for signal transduction and other functions
of membrane proteins. It has been suggested that lateral diffusion
of lipids in cellular membranes regulates the formation and
dispersion of these lipids nanodomains and thus the functions of
membrane proteins (Simons and Toomre, 2000; Sprong et al.,
2001; Lingwood and Simons, 2010). Therefore, characterizing
the dynamics of lipids in membranes and elucidation of the
mechanisms that regulate this motion are crucial. In addition,
recent evidence has demonstrated that the plasma membrane is
compartmentalized by actin-dependent membrane protein fences
(Kusumi et al., 2011). When proteins or lipids are imaged at high
acquisition rates, they display a non-Brownian diffusion pattern
owing to the obstacles that temporarily restrict their movements
(Ritchie et al., 2005). A variety of optical techniques (and
variations of the theme) have been used to determine the
movement of lipids and proteins in the plane of the membrane,
including FRAP, FCS and SPT. FRAP and FCS approaches
are frequently used to measure the long-range diffusion and
fluctuations of a population of fluorescent molecules,
respectively, whereas SPT is a robust method used to analyze
the mobility of individual lipids (or proteins) in the PM of living
cells and in model membranes. For these techniques, fluorophorelabeled lipids, toxin domains and genetically encoded proteins are
suitable as lipid probes. Below, we will highlight examples of the
use of each of these techniques and describe how they can
generate complementary data sets.
Fig. 2. High-resolution imaging of lipids using EM, PALM and STED.
(A) EM images of localization of PtdIns3P labeled with GST–26FYVE are
shown. An ultrathin frozen section of fixed baby hamster kidney cells was
incubated with GST–26FYVE. After washing and fixation, samples were
incubated with anti-GST antibody and then 10-nm gold particles conjugated
to protein A (shown by arrows). Clear cytoplasmic coats of early endosomes
are shown by arrowheads (left). The multi-vesicular bodies show labeling of
GST–26FYVE concentrated on the internal membranes (right). Scale bars:
200 nm. Images reproduced from Gillooly et al., 2000 with permission from
EMBO J. (B) SM-enriched domains in the outer leaflets of PM were
visualized by PALM. HeLa cells were incubated with recombinant Dronpa–Nterminal truncated Lysenin at room temperature for 10–60 min, washed
quickly, fixed with 4% formaldehyde in PBS and observed with PALM
microscopy. Expansion of the region indicated by the white box is shown on
the right side. Scale bars: 500 nm. Images reproduced from Mizuno et al.,
2011 with permission from The Royal Society of Chemistry. (C) Confocal
(left) and corresponding STED (right) images of PtdIns(4,5)P2 in the cytosolic
leaflets of PM are shown. PtdIns(4,5)P2-enriched domains in the cytosolic
leaflets of PM are visualized with high resolution by STED. The resolution is
increased in the STED image compared to the corresponding confocal
image. Membrane sheets of PC12 cells were prepared by rupturing the cells
with probe sonication as described previously (Sieber et al., 2006). Then, the
membrane sheets were immunostained with a monoclonal anti-PtdIns(4,5)P2
antibody (2C11) and a secondary antibody labeled with Alexa Fluor 488, and
observed with STED microscopy. Scale bars: 5 mm. Images reproduced from
van den Bogaart et al., 2011 with permission from Nature.
4806
FRAP analysis with lipid probes can be performed by standard laser
scanning or spinning-disk confocal microscopy to irreversibly bleach
the fluorescence of probes in a region of interest (Axelrod et al., 1976;
Corbett-Nelson et al., 2006; Hammond et al., 2009b; Kay et al., 2012).
In some cases, total internal reflection fluorescence (TIRF)
microscopy is also used for FRAP analysis (Axelrod, 2001;
Hammond et al., 2009b). The region that has been photo-bleached
is then monitored by time-lapse microscopy to determine the kinetics
of the recovery of the fluorescent signal (Lippincott-Schwartz et al.,
2003; Lippincott-Schwartz and Patterson, 2003). As several of the
lipid probes used for FRAP have a high rate of association and
disassociation, additional post-acquisition analysis must be performed
to elucidate the actual diffusion rate of the lipid under investigation
(Fig. 3A; for more details, see Hammond et al., 2009b).
Recent studies have investigated the lateral diffusability of
PtdIns(4,5)P2 in the plasma membrane by FRAP analysis
using both protein-based probes and fluorophore-labeled
lipids (Hammond et al., 2009b; Golebiewska et al., 2011).
Plasmalemmal PtdIns(4,5)P2 is a spatial signal and contributes to
the identity of the PM through the recruitment of effectors. By
determining the diffusion rates of PtdIns(4,5)P2 in the membrane,
researchers can estimate the area an effector protein can survey
while it is bound to PtdIns(4,5)P2 if the rate of dissociation is
known. Using the GFP–PLCd-PH and BODIPY–PtdIns(4,5)P2,
the lateral diffusion (D) was found to be ,1 mm2/s. The
disassociation time (t) of the GFP–PLCd-PH from this lipid is
estimated to be 2.4 s (Hammond et al., 2009b). Therefore, the
distance traveled by the protein while bound to the lipid can be
estimated by the equation !(26D6t) (Teruel and Meyer, 2000).
This suggests that PLCd or other effectors with comparable
affinity for this lipid can potentially travel 1–3 mm in the plane of
the membrane before a significant fraction of the protein
Journal of Cell Science
FRAP
COMMENTARY
Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
disassociates (Hammond et al., 2009b) This ability to diffuse in
the plane of the membrane is likely an important contributor to
the formation of protein–protein interactions and macromolecular
complexes (Fig. 3B; Golebiewska et al., 2011). The study of
Hammond et al., did not characterize the presence of a diffusion
barrier for PtdIns(4,5)P2, but the study using the BODIPY–
PtdIns(4,5)P2 demonstrated that this lipid does not diffuse out of
the forming phagosome, suggesting that the there is a diffusion
barrier at the tips of the pseudopods that drives phagosome
formation (Golebiewska et al., 2011).
A similar study examined the dynamics of PtdSer using
TopFluor–PtdSer and GFP–Lact-C2 using FRAP in HeLa cells
(Kay et al., 2012). The mean diffusion coefficient of expressed
GFP–Lact-C2 in the PM of HeLa cells was determined to be
0.33 mm2/s compared to 0.49 mm2/s for the TopFluor–PtdSer
(Kay et al., 2012). This value is substantially lower than the
diffusion coefficients for the PtdIns(4,5)P2 probe (GFP–PLCdPH), suggesting that PtdSer might interact with other lipids or
proteins to limit its diffusion. Support of this concept is provided
by the fact that only 43% of the TopFluor–PtdSer is mobile in the
FRAP experiments. Taken together, the results suggest that there
is a fraction of PtdSer that is either immobile or greatly restricted
in its lateral mobility. One possibility is that the PtdSer clusters
observed by EM might in fact be sequestered PtdSer molecules
that are confined by interactions with cholesterol and/or protein
complexes (Fairn et al., 2011b; Kay et al., 2012).
FCS
FCS encompasses many related techniques to analyze
fluctuations in fluorescence for populations or individual
A
Pre-bleaching
(ii)
Bleaching
(iii)
Recovery
Medium or
cytosol
Membrane
Lipid probe
(lipid−binding domain)
B
(i)
Pre-bleaching
Diffusion
Bleached lipid probe
Association/dissociation
(ii)
Bleaching
(iii)
Recovery
Medium or
cytosol
Membrane
C
GFP−Lact-C2
Raw data
Diffusion
Bleached lipid probe
Classified tracks
(iii)
(iii)
Fraction of tracks
Lipid probe
(fluorophore labeled lipids)
Fig. 3. Dynamics of lipids analyzed by
FRAP and SPT. (A,B) Schematic diagram
of the FRAP analysis. The fundamental
differences of the mode of recovery for
these types of lipid probes are shown
(Hammond et al., 2009b; Kay et al., 2012).
(A) Following the photobleaching of the
lipid-binding domains that are produced in
the cytosol, recovery will occur by two
means; diffusion of probes while they are
bound to their lipid ligand (a red two-way
arrow in iii) and dissociation of bleached
probes and association of unbleached
probe from the cytosolic pool (a blue twoway arrow in iii). (B) Using fluorophorelabeled lipids, probes in both leaflets are
photo-bleached (ii) before fluorescence is
recovered through the diffusion of the lipids
only (a red two-way arrow in iii). (C) SPT
analysis of GFP–Lact-C2 in the cytosolic
leaflets of PM. GFP–Lact-C2 was
transfected to HeLa cells and SPT using
TIRF microscopy was performed. Raw
images and data are reproduced from Kay
et al., 2012 with permission from Molecular
Biology of the Cell. Representative image
sequences (i) and track classification (ii) of
GFP–LactC2 on the inner leaflet of the PM
of HeLa cells are shown. Scale bar: 2 mm.
(iii) The graph shows the breakdown of the
tracks illustrated in (i).
1.0
0.8
Journal of Cell Science
(i)
0.6
0.4
0.2
n
Co
fin
ed
Fr
ee
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Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
SPT
SPT techniques have been used largely in the field of membrane
biophysics to determine the lateral diffusion of lipids and proteins
in both PM and supported bilayers (Saxton and Jacobson, 1997;
Martin et al., 2002). TIRF microscopy can be combined with
SPT, and this system is particularly sensitive for measuring the
dynamics of lipid probes at the surface of either model
membranes (in vitro) or living cells (in vivo) (Knight and
Falke, 2009; Kay et al., 2012). Although FRAP is able to
determine the average dynamics of hundreds or thousands of
molecules, SPT measures the dynamics of individual molecules
in considerably shorter time intervals and with greater precision.
Thus, lipid subpopulations that cannot be identified using FRAP
can be categorized by SPT analysis (Saxton and Jacobson, 1997).
In SPT, time-lapse sequences of images of probes are taken with
fluorescence microscopy and analyzed with a variety of
algorisms. Typically, the position and intensity of a particle is
determined by detecting local maximum intensities and then
fitting Gaussian curves that approximate the two-dimensional
point spread function of the microscope (Jaqaman et al., 2008;
Flannagan et al., 2010; Jaqaman et al., 2011). Using SPT, a study
compared the mobility of two PtdSer probes, TopFluor–PtdSer
and GFP–Lact-C2, in the PM (Kay et al., 2012). Interestingly,
only 29% of the TopFluor–PtdSer is freely diffusible with the
remaining 71% being confined to smaller areas that have an
average radius of 111 nm. Whereas the analysis of the GFP–LactC2 probe showed that 78% of the GFP–Lact-C2 displays free
diffusion and the remaining 22% are confined within an average
area of 360 nm (Fig. 3C) (Kay et al., 2012). Clearly the two
different PtdSer probes are generating opposing data, but the
question is why? Like all translocation-based protein domain
probes, Lact-C2 can only bind to the unengaged head group and
therefore ‘available’ PtdSer. Therefore, the results would suggest
that the majority (,80%) of the PtdSer that is unbound or
unshielded by proteins is freely diffusible. By contrast, the acylchain-modified TopFluor–PtdSer can be identified and tracked
regardless of it binding to proteins or being in complexes such as
caveolae. The TopFluor–PtdSer result is consistent with a large
percentage of the plasmalemmal PtdSer being bound or occupied
by proteins or membrane domains. This reinforces the notion that
researchers must be aware of the nature of the probe being used to
have a full appreciation of the results. These differences also
demonstrate the utility of using multiple probes for a specific
lipid to gain a better understanding of its behavior in vivo.
Similar experiments have been used to follow the dynamics
of PtdIns(3,4,5)P3 on supported lipid bilayers (Knight and
Falke, 2009). Here, PtdIns(3,4,5)P3 was monitored using an
A
B
C
Filipin
Topfluor−Cholesterol
Dehydroergosterol
Fig. 4. Intracellular distribution of cholesterol visualized by established cholesterol probes. (A) Filipin allows visualization of cholesterol in the PM,
recycling endosomes and other intracellular vesicles. For filipin staining, CHO cells were fixed with 3.7% formaldehyde in PBS, then cells were treated with
0.5 mg/ml filipin at 4˚C for 16 h. Images were acquired by laser scanning confocal with a 405-nm laser. (B) TopFluor–cholesterol accumulates in intracellular
vesicles after incubation at 37˚C. A total of 10 nmol/ml TopFluor–cholesterol in PBS containing 1% BSA was loaded to CHO cells for 10 min on ice. After washing
with PBS, cells were cultured at 37˚C for 10 min, then cells were fixed with 3.7% formaldehyde in PBS. Images were acquired using confocal microscopy.
(C) DHE accumulates in PM and recycling endosomes. TRVb1 cells, a CHO cell line that expresses human transferrin receptor, were labeled with a DHE–
methyl-b-cyclodextrin complex for 1 min, washed and chased at 37˚C for 30 min, then images were obtained by live-cell imaging (Mondal et al., 2009). Images in
A and B were obtained with laser scanning confocal microscopy. The image in C was obtained with a non-confocal inverted fluorescence microscopy optimized
for DHE imaging as described by Mukherjee et al., 1998. Reproduced from Mondal et al., 2009 with permission from Molecular Biology of the Cell. Scale bars:
10 mm.
4808
Journal of Cell Science
labeled molecules diffusing into and out of an excitation region.
This excitation region can be a focal-spot or line-scan that is
monitored over time using either single- or two-photon excitation.
(Fahey et al., 1977; Korlach et al., 1999; Schwille et al., 1999;
Haustein and Schwille, 2007). Line-scan FCS has been used to
examine both the diffusion of BODIPY–cholesterol in liquid
ordered and disordered domains in phase-separating model
membranes. Using a variety of phosphatidylcholine (PtdCho),
sphingomyelin and cholesterol model membranes, a study found
that the diffusion coefficient was approximately an order of
magnitude higher in liquid-disordered regions compared to the
liquid-ordered regions (Ries et al., 2009). When the same
experiment was conducted with HEK cells, the BODIPY–
cholesterol had a diffusion coefficient of 0.33 mm2/s, which is
comparable to that of PtdSer. The dynamics of BODIPY–
PtdIns(4,5)P2 in the inner leaflets of PM and in forming
phagosomes has also been analyzed with FCS and was found to
have a diffusion coefficient of ,1 mm2/s, in both forming
phagosomes and the PM of J774 macrophages (Golebiewska
et al., 2008; Golebiewska et al., 2011).
FCS analysis of lipids can be influenced by a number of
factors, including the relative amount of cytosol in the region of
interest. For example, FCS analysis detected both a fast and slow
population of molecules in the study using the TopFluor–PtdSer
and GFP–Lact-C2 probes in the PM of HeLa cells (Kay et al.,
2012). The diffusion coefficient of the slow fraction likely
reflects the population of TopFluor–PtdSer in the membrane and
the GFP–Lact-C2 bound to PtdSer, respectively. By contrast,
the fast-moving fraction of these probes is likely a soluble
degradation product of the TopFluor–PtdSer and the unbound
(cytosolic) GFP–Lact-C2. Thus, understanding the nature of the
probe used can assist in the interpretation of the FCS results.
Alexa-Fluor-555-conjugated GST–GRP1-PH in order to
determine both the diffusion coefficient for the PtdIns(3,4,5)P3
and dissociation rate of the probe (Knight and Falke, 2009). This
type of in vitro analysis will allow researchers to move to in vivo
systems and gain a better understanding of PtdIns(3,4,5)P3
signaling. Overall, SPT analysis is a powerful tool to observe
lipid (and protein) dynamics and to obtain new insights into
the molecular mechanisms of lipid–lipid and/or lipid–protein
interactions in membranes.
Future perspectives
In this Commentary, we have highlighted established probes for
a variety of lipids (Table 1); however, lipids probes are not
currently available for all classes of lipids. For instance, proteinbased probes for visualizing intracellular phosphatidylcholine and
phsophatidylethanolamine (PtdEtn) have not yet been identified.
It has been reported that the short polypeptides duramycin and
cinnamycin specifically bind to PtdEtn (Navarro et al., 1985;
Choung et al., 1988), and cinnamycin has been used visualize
PtdEtn in the outer leaflets of PM during cytokinesis and
apoptosis (Emoto et al., 1996; Makino et al., 2003). However,
these molecules cause cell lysis at high concentrations.
Additionally, fixation and/or permeabilization are necessary for
the compounds to gain access to intracellular compartments
making them less than ideal. In addition, although there are
many well-established probes for other phosphoinositides,
specific probes for phosphatidylinositol 3,5-bisphosphate
[PtdIns(3,5)P2] and phosphatidylinositol 5-phosphate [PtdIns5P]
have been lacking. However, the cytosolic N-terminal polybasic
domain of TRPML1 (ML1N) has recently been reported to
specifically bind to PtdIns(3,5)P2 (Li et al., 2013). This domain
awaits further characterization but appears to be suitable to
analyze the intracellular localization of PtdIns(3,5)P2 using
light microscopy. Clearly, the isolation of probes with greater
specificity and/or affinity will provide more accurate information
about lipids or specific pools of lipids. A prime example of this is
the characterization of a new phosphatidylinositol 4-phosphate
[PtdIns4P] probe, P4M that demonstrates the presence of this
lipid in late endosomes/lysosomes (Hammond et al., 2014).
Cholesterol is one of the most abundant lipid molecules in the
PM, the cell and within the body. Recent studies have established
a concept that cholesterol forms membrane nanodomains (lipid
rafts) in the outer leaflets of PM (Lindwood and Simons, 2010).
Lipid rafts represent a scaffold where specific proteins assemble
and play important roles in many cells functions such as signal
transduction and virus infection (Simons and Toomre, 2000;
Simons and Ehehalt, 2002). Several probes for cholesterol have
been established to visualize intracellular cholesterol (Fig. 4, see
also Box 1). Although these cholesterol probes can label
cholesterol in cellular membranes, the big problem is that these
probes cannot distinguish cholesterol in the outer leaflets and in
the inner leaflets of PM (Ohno-Iwashita et al., 2010a; see also
Box 1). Thus, design of new cholesterol probes that can
specifically visualize cholesterol in the inner leaflets of PM
would be an important future work.
Finally, recent technological advances have produced a variety
of super-resolution microscopic techniques, for example,
structured illumination microscopy (SIM), STED, PALM and
STORM. However, these still have limitations that need to be
overcome for their widespread use in the imaging of lipids
(Galbraith and Galbraith, 2011). For instance, high image
resolution often trades off with acquisition speed, which is
Journal of Cell Science (2014) 127, 4801–4812 doi:10.1242/jcs.150524
Box 1. Technical challenges for the visualization of
intracellular cholesterol
Cholesterol is an essential structural component of cellular
membranes and has pivotal roles in signal transductions and
membrane trafficking (Ikonen, 2008). Established methods for
visualization of intracellular cholesterol are discussed below.
Filipin
The antibiotic filipin is an intrinsically fluorescent polyene
compound, which specifically binds to free cholesterol (Börnig
and Geyer, 1974). Filipin is routinely used for visualization of free
cholesterol in fixed cells. In CHO cells, PM and endosomes are
stained by filipin in CHO cells (Fig. 4A). Notably, filipin can stain
free cholesterol in all of the intracellular membrane compartments
because of its membrane permeability (Miller, 1984; Ohno-Iwashita
et al., 2010b). Thus, filipin staining is not an appropriate method for
live-cell imaging. In addition, filipin photobleaches very quickly.
Thus, it can be challenging to obtain high-quality images with a
standard confocal microscopy that typically use 405 nm lasers.
TopFluor–cholesterol and BODIPY–cholesterol
TopFluor–cholesterol is a fluorophore-labeled cholesterol (Li et al.,
2006; Hölttä-Vuori et al., 2008). TopFluor–cholesterol can be easily
loaded to cells with BSA and mainly localizes in endosomes in CHO
cells (Fig. 4B). One concern is that the addition of the fluorophore
might alter the properties and dynamics of cholesterol and thus might
not accurately mimic endogenous cholesterol (Solanko et al., 2013).
In addition, TopFluor–cholesterol cannot distinguish the cholesterol in
the cytosolic leaflets and luminal leaflets cellular membranes.
Dehydroergosterol
Dehydroergosterol (DHE) is a naturally fluorescent sterol and a
cholesterol analog. In cells loaded with DHE, most of the fluorescent
signal accumulates in the recycling endosomes and to a lesser extent
in the PM in CHO cells (Fig. 4C). One advantage is that DHE is
suitable for live-cell imaging experiments, but overall it is weakly
fluorescent and is typically used with a modified wide-field microscope.
Previous studies using fluorescence quenchers have demonstrated
that the majority of the DHE is localized within the inner leaflet of the
PM (Mondal et al., 2009). This result was surprising considering the
belief that most cholesterol in the PM is localized to the exofacial
leaflet. It is unclear if the observations using DHE are true for
cholesterol (Hyslop et al., 1990). However, it should be noted that the
vast majority of cholesterol could be replaced by DHE without the loss
of cell viability (Hao et al., 2002; Wüstner, 2007; Mondal et al., 2009).
important as lipids and proteins can potentially move faster than
the speed with which super-resolution microscopes capture
images. To overcome these issues, we will need to use photoswitchable probes and small-molecule fluorophores for superresolution microscopes, in addition to technical improvement for
temporal-spatial resolution (Fernández-Suárez and Ting, 2008).
As illustrated throughout this Commentary, BODIPY has been
used extensively to generate fluorophore conjugated lipids.
However, BODIPY has relatively poor photo-stability compared
to several other fluorescent dyes. An alternative dye that could
become more popular is ATTO647N, which is routinely used in
STED microscopy. ATTO647N has been used to label either the
head group or acyl chain of a variety of lipids, such as PtdEtn,
sphingomyelin and the ganglioside GM1 (Eggeling et al., 2009).
Indeed, the dynamic behavior of various lipids in the PM of living
cells has already been measured in greater detail with the
combined use of FCS and STED (Eggeling et al., 2009; Mueller
et al., 2011), which enhanced resolution. We therefore believe
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COMMENTARY
that in the near future analysis of lipid dynamics by FCS and SPT
combined with super-resolution microscopy will provide new
insights into the cell biology of lipids and membranes.
Acknowledgements
We thank Sergio Grinstein for helpful discussions and Takashi Hirama (both
Hospital for Sick Children, Toronto, Canada) for technical advice regarding the
use of the BODIPY–LacCer. We apologize to all colleagues whose work was not
cited due to space restrictions.
Competing interests
The authors declare no competing interests.
Funding
G.D.F. is a recipient of a New Investigator Salary Award from Canadian Institutes
of Health Research (CIHR) and an Ontario Early Research Award. Work in our
laboratory is supported by St. Michael’s Hospital New Investigator Start-up Fund,
an operating grant from CIHR and a Discovery Grant from the Natural Sciences
and Engineering Research Council of Canada.
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