Biochem. J. (2010) 430, 21–37 (Printed in Great Britain)
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doi:10.1042/BJ20100372
REVIEW ARTICLE
Tracking the green invaders: advances in imaging virus infection in plants
Jens TILSNER* and Karl J. OPARKA†1
*Plant Pathology Department, Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, U.K., and †Institute of Molecular Plant Sciences, University of Edinburgh, Mayfield
Road, Edinburgh EH9 3JH, U.K.
Bioimaging contributes significantly to our understanding of
plant virus infections. In the present review, we describe
technical advances that enable imaging of the infection process
at previously unobtainable levels. We highlight how such new
advances in subcellular imaging are contributing to a detailed
dissection of all stages of the viral infection process. Specifically,
we focus on: (i) the increasingly detailed localizations of
viral proteins enabled by a diversifying palette of cellular
markers; (ii) approaches using fluorescence microscopy for the
functional analysis of proteins in vivo; (iii) the imaging of
INTRODUCTION
In order to infect a host cell and to spread systemically, viral
pathogens have to usurp, manipulate and overwhelm numerous
biological processes within cells. They often achieve this with
a minimal set of multifunctional virus-encoded proteins, as well
as through the recruitment of host factors [1–5]. Most of these
‘hijacked’ processes are compartmentalized within the host cell,
and thus the virus and its gene products become localized to
specific subcellular compartments during the appropriate stages
of infection, often in a highly dynamic manner. The ability
to visualize these events and their ensuing effect on the host
cell have greatly enhanced our understanding of the infection
process.
Since the 1950s, EM (electron microscopy), later coupled
to immunogold detection of specific proteins and RNA, has
been the mainstay of virological approaches, but is restricted
to static snapshots of the infection process. Furthermore, the
limited accessibility of antigenic epitopes in samples embedded
and sectioned for EM makes it difficult to detect less-abundant
proteins or to quantitatively assign visible structures to specific
biomolecules [6]. The discovery of FPs (fluorescent proteins) reintroduced LM (light microscopy) as a major tool for cell biology
in the early 1990s, and today fluorescence microscopy is one
of the cornerstones of plant virology. Initially, virus-expressed
FPs were simply used to monitor the spread of infection [7,8],
but plant virologists quickly started to employ them as fusions
to virus-encoded proteins. Nowadays, fluorescence microscopy
is used increasingly to permit not just localization, but also
viral RNAs; (iv) methods that bridge the gap between optical
and electron microscopy; and (v) methods that are blurring the
distinction between imaging and structural biology. We describe
the advantages and disadvantages of such techniques and place
them in the broader perspective of their utility in analysing plant
virus infection.
Key words: correlative microscopy, fluorescent protein, in vivo
interaction, membrane topology, RNA imaging, super-resolution.
functional analysis of proteins in vivo, and new approaches
have allowed optical-based microscopy to break the diffraction
barrier, long held as a fixed limit to resolution [9–11]. At the
same time, EM has also made important advances, in particular
through improved sample preservation using high-pressure
freezing/freeze substitution [12] and the extension into 3D (threedimensional) imaging using ET (electron tomography) [13,14].
Consequently, correlative approaches that combine LM and EM
are currently being developed in many laboratories [15,16]. CryoEM tomography and AFM (atomic force microscopy) are starting
to blur the boundary between imaging and structural biology,
and are opening the door to a truly molecular understanding of
the viral infection process. In the present review, we summarize
some of these recent developments with a particular focus on the
challenges and advantages that are specific to imaging plant virus
infection.
ADVANCEMENT OF PROTEIN LOCALIZATION STUDIES
Since the first use of GFP (green FP) as a reporter of virus infection
in plant cells [7], the palette of autofluorescent proteins available
for localization studies has increased exponentially through the
discovery of new natural source proteins and the engineering
of new varieties. Generally, new FPs are selected to increase the
number of spectral variants that can be separated in co-localization
experiments, and to overcome problems with photostability,
brightness, oligomerization and fusion tolerance [17,18]. For plant
virus studies, however, brightness and photostability are often
less important considerations than spectral properties because
Abbreviations used: 3D, three-dimensional; AFM, atomic force microscopy; BiFC, bimolecular fluorescence complementation; CFP, cyan fluorescent
protein; CLEM, correlative light and electron microscopy; COP, coatamer protein; CP, capsid protein; DAB, diaminobenzidine; dsRNA, double-stranded
RNA; EM, electron microscopy; ER, endoplasmic reticulum; ET, electron tomography; EYFP, enhanced yellow fluorescent protein; FlAsH, fluorescein
arsenical helix binder; FLIM, fluorescence lifetime imaging; FLIP, fluorescence loss in photobleaching; FP, fluorescent protein; FPC, C-terminal fragment
of split FP; FPN, N-terminal fragment of split FP; FRAP, fluorescence recovery after photobleaching; FRET, fluorescence resonance energy transfer;
GFP, green fluorescent protein; LM, light microscopy; MP, movement protein; N, nucleocapsid; ORF, open reading frame; PA, photoactivatable; PALM,
photoactivation localization microscopy; pc-FP, photoconvertible fluorescent protein; PD, plasmodesma(ta); PSF, point-spread function; PUMHD, Pumilio
homology domain; PVX, potato virus X; Qdot, quantum-dot; ReAsH, resorufin arsenical helix binder; RFP, red fluorescent protein; RNP, ribonucleoprotein
particle; ROI, region of interest; SIM, structured illumination microscopy; STED, stimulated emission depletion; STORM, stochastic optical reconstruction
microscopy; TGB, triple gene block; TMV, tobacco mosaic virus; VRC, viral replication centre; vRNA, viral RNA; YFP, yellow fluorescent protein.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2010 Biochemical Society
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J. Tilsner and K. J. Oparka
transient expression from agrobacteria, particle bombardment
or viral genomes is the most common experimental approach.
As a result, researchers usually deal with problems related to
overexpression rather than detectability or dimness, and FPs
for plant virus studies can simply be chosen based on their
spectral separation. Because virus-expressed FP constructs can
accumulate at extremely high levels, sequential image acquisition
is preferable for co-localizations, at least as a control. Several sets
of expression vectors are now available that enable fast generation
of multiple FP-fusions from a single clone of the gene of interest
based on Invitrogen’s proprietary Gateway in vitro recombination
system [19–31], and additional vectors using more recently
developed FP variants are continuously being developed [30,31].
As knowledge of cell biology increases, so does the number
of available fluorescent markers for subcellular compartments
and transport pathways [32,33], and plant virologists are taking
advantage of this to determine the location of viral proteins in
detail, and to follow their trafficking pathways throughout the
cell. For instance, the 6K protein of potyviruses, which functions
to establish viral replication sites, was known previously to target
the ER (endoplasmic reticulum) [34]. Wei and Wang [35] have
shown that the specific location of the 6K protein is the ER exit
site, where 6K accumulation depends on both the COP (coatamer
protein) I and COPII vesicle trafficking machineries, and that the
6K-containing membrane compartments travel onwards through
the secretory pathway to chloroplast membranes where they
accumulate to form the main replication sites [36] (Figures 1a and
1b). In another study, Taliansky and co-workers used a variety of
markers of subnuclear bodies to follow the path of the ORF3 (open
reading frame 3) protein of the umbravirus groundnut rosette
virus, as it enters the nucleus [37,38]. Here it interacts with Cajal
bodies, fuses them with the nucleolus and recruits the nucleolar
RNA-binding protein fibrillarin for export into the cytoplasm and
incorporation into viral movement complexes (see Figure 3c).
Considerations specific to localizing viral proteins
As experiments designed to track the subcellular localizations
of virus-encoded proteins become more sophisticated, careful
interpretation of the results also becomes more critical. Previous
reviews have covered such issues in a more general context [39–
41], but for studies of viral proteins a number of additional
considerations apply (Figure 2). Three important factors are:
(i) expression levels; (ii) potential interaction partners during
infection (nucleic acids as well as proteins); and (iii) steric
interference of the FP tag with protein localization and function.
For endogenous plant proteins, the ‘gold standard’ control for
all three criteria is to complement a knockout mutation of the gene
of interest with the FP-fusion expressed from the gene’s native
promoter. The equivalent to this for a virus-encoded protein would
be to replace the original ORF with the FP-fused gene in the viral
genome. However, this is often not possible. Viral genomes have
been selected for extremely economic use of genomic capacity.
Often ORFs overlap, or are expressed as polyproteins that are
cleaved in tightly regulated processing pathways. The gene of
interest may also overlap with (subgenomic) promoters or other
regulatory sequence elements [42,43]. Viral genome sizes are
often limited by virion packaging mechanisms, constraining FP
insertions, and in RNA viruses, high recombination rates can
quickly eliminate insertions. With the additional potential for FPfusions to disrupt the (multi)functionality of viral proteins, it is
not surprising that approaches in which the FP-fusion functionally
replaces the wild-type gene in the viral genome (e.g. [44,45]) are
the exception rather than the rule. Instead, various compromises
have been used such as insertion under the control of alternative
c The Authors Journal compilation c 2010 Biochemical Society
viral promoters (e.g. [7,34,46]), self-cleaving FP-fusions using a
linker derived from the 2A protein of foot and mouth disease
virus to release sufficient native viral protein for functional
complementation [47] or expression from viral satellite genomes
(e.g. [48]). Such viral expression strategies, together with
agroinfiltration and particle bombardment, all comprise transient
expression assays, which are convenient and produce fast results.
They are also somewhat appropriate, as viral proteins in most
cases are only ‘transiently’ expressed during a natural infection,
and often to very high levels. Additionally, transient expression
can be advantageous compared with transgenic plants stably expressing viral proteins, as the latter are often resistant to infection.
Where localization studies differ most significantly between
viral and plant proteins is the nature of the virus as a selfcontained ‘closed’ system. Redundancy between functionally
similar genes or lack of mutant phenotypes is not usually a
problem in virus systems. On the other hand, expression of
individual viral proteins can never be representative for the context
of the actual infection and, by whatever means, localizations
should be studied in infected tissue [49]. Most likely, the viral
protein of interest will be affected by interactions with other
viral or infection-specific factors. If non-viral expression is
the only option, 35S-driven viral replicons can be a useful
tool to spatially and temporally control co-introduction of the
fluorescent reporter with the virus. This can also facilitate studies
of movement-deficient or replication-attenuated viral mutants.
However, some viral proteins may only function (and localize
correctly) when expressed in cis. But the ability to compare
localizations between uninfected and infected tissue provides an
additional tool to dissect these interactions and functions, similar
to genetic approaches using knockout mutations (Figure 2).
Although high transient expression levels are not problematic
themselves, localizations should consider the timing and
expression levels during infection. For example, plant virus MPs
(movement proteins) are often expressed early and at low levels,
whereas CPs (capsid proteins) are typically expressed later and
at very high levels. Therefore prolonged overexpression of an
MP–FP may need to be interpreted with more caution than a
CP–FP fusion. After particle bombardment, FP fluorescence can
often be observed after a few hours, and in agroinfiltration and
viral expression systems, imaging may be possible as early as
1 day after inoculation. In view of the fact that many viruses
replicate and move intercellularly within less than 1 day [50–
53], but may continue to accumulate progeny virions until the
host cell’s resources are exhausted, the localization of a viral
protein should be studied as early as possible as well as at later
stages to observe changes in localization and function during
the infection cycle. Within a viral lesion, the differences in
localizations between cells at the leading edge (= early) and the
centre (= late) of an infection site offer a valuable reference in this
regard. Owing to the maturation time of FPs and requirements for
replication or subgenomic RNA production, virus-expressed FPfusions are generally unsuitable to study the earliest events during
the infection process. Virus-independent transient expression can
be a way to circumvent this problem, as the protein of interest
can be present prior to infection, for instance by allowing a virus
infection to enter an area already expressing the FP-fusion of
interest. Again, the potential requirement for expression in cis
needs to be kept in mind.
In transient experiments using non-native expression, relative
ratios of potential interaction partners also have to be considered.
For example, the overlapping ORFs of the second and third ‘triple
gene block’ MPs of different viruses are expressed from a single
subgenomic messenger RNA, with the downstream TGB3 (triple
gene block 3) ORF translated by leaky ribosome scanning after
Imaging plant virus infections
Figure 1
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Various applications of live-cell fluorescence microscopy suitable for plant virus imaging
I. Detailed localizations within the context of the host cell. (a) Potyviral 6K replication protein induces ER-derived vesicles that co-localize with the ER-to-Golgi exit site marker Sar1 (i, ii). A
dominant-negative Sar1 mutation abolishes 6K vesicle formation (iv, v). At later stages, the 6K-induced vesicles transfer to the chloroplast envelope (iii), the putative viral replication site. Transport
depends on the actin cytoskeleton, shown by reduced plastid labelling after overexpression of a dominant-negative fragment of the motor protein myosin IX-K (vi) (reproduced from [35,36] with
permission from American Society for Microbiology). II. Study of protein dynamics. (b) FRAP of PD-localized TMV MP shows that MP is delivered actively to PD: the metabolic inhibitor azide (lower
panels) blocks fluorescence recovery (modified from [83] with permission from John Wiley and Sons). (c) Use of the green-to-red photoswitching protein Kaede to localize translation sites. At time
0 min, a patch of Kaede-labelled potassium channel Kv1.1 in a neuronal dendrite (outlined) is converted into red. Within 60 min, newly synthesized green Kv1.1-Kaede has accumulated, which is
again converted into red at time 60 min. Only green fluorescence is shown (modified from [89]. Reprinted with permission from AAAS). III. Detection of protein–protein interactions. (d) FLIM–FRET
of tomato spotted wilt virus replication proteins. N protein co-localizes with both GP and Gn, but only interacts directly with GP, as shown by reduced CFP fluorescence lifetime (co-localization
c 2009, with permission from Elsevier). IV. Topology analysis of integral membrane proteins. (e) BiFC-based membrane
and FLIM–FRET images show different cells) (modified from [189] c The Authors Journal compilation c 2010 Biochemical Society
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Figure 2
J. Tilsner and K. J. Oparka
Complementing approaches for studying the subcellular locations of plant viral FP-fusion proteins in the context of infection
initiation at the upstream TGB2 ORF. This results in an excess
of TGB2 protein over TGB3 [54]. Recently, Lim et al. [48,55]
have shown that the TGB2/TGB3 ratio profoundly influences the
TGB3-dependent localization of the TGB2 protein of barley stripe
mosaic virus.
Finally, the choice of host system needs to be considered.
Compared with animal virologists, researchers working on plant
viruses are in the advantageous position of having both intact
plants and isolated cells for infection, but both come with
certain limitations. In some cases, for instance with fusionintolerant viral proteins (often replicases) or specific RNAs
(discussed below), immunofluorescence is a better option than
FP-fusions. However, to make intracellular epitopes accessible
to antibody labelling, fixation and membrane permeabilization
are usually insufficient in plant tissues due to the presence
of the cell wall, requiring additional embedding and sectioning.
The use of protoplasts can overcome this limitation and offer the
additional benefit of permitting synchronized infection of an
entire culture, i.e. defined infection timing [56]. Protoplasts
are also more accessible for invasive delivery methods such as
membrane permeabilization or microinjection that may open new
avenues in the investigation of early infection events (see below).
However, even when prepared from fresh tissue rather than cell
culture, they represent profoundly altered non-native host cells.
Aditionally, many membranous organelles, which are important
in viral processes such as replication and movement, are strongly
perturbed by fixation and embedding. For imaging, especially in
vivo, intact tissue should be the preferred choice, with protoplast
systems reserved for more specialized experimental problems.
LABELLING FUSION-INTOLERANT VIRAL PROTEINS WITH SMALL
FLUORESCENT TAGS
Besides expression level and context, the most important factor
influencing the accuracy of fluorescence-based localizations is
the effect of the fluorescent tag on the protein of interest. FPs
may mask functional domains, interfere with interactions with
other macromolecules, or have a stabilizing or destabilizing effect.
Simply increasing protein size by adding a ∼ 27 kDa GFP moiety
topology analysis of potato mop-top virus TGB2 protein. YFPN (yellow FPN) fragment fused to the N- or C-termini of TGB2 results in BiFC with cytoplasmic YFPC (yellow FPC) (i, iii), whereas
YFPC introduced into the hydrophilic loop of TGB2 complements ER-luminal YFPN (ii), leading to the topology models on the right. (modified from [62] with permission from John Wiley and
Sons). (f) Redox-sensitive GFP-based topology analysis. roGFP is more strongly excited by 405 nm illumination in an oxidizing environment such as the ER lumen. Fusion of roGFP to the ends of
transmembrane proteins results in distinct 405/488 ratios depending on the cellular compartment which the protein terminus is facing. Reproduced from [121] with permission from John Wiley and
Sons. V. RNA imaging. (g) PUMHD-BiFC vRNA imaging. In the VRC of TMV (left), RNA is localized to punctate ‘hot spots’, whereas in the PVX VRC (right), it is arranged in circular ‘whorls’. Red
fluorescence from virally expressed RFPs was used as infection markers [105]. Scale bars: (a, panels i, ii, iv and v), 12 μm; (a, panels iii and vi), 8 μm; (b), (d) and (e), 5 μm; (c), 2 μm; (f) and
(g), 10 μm.
c The Authors Journal compilation c 2010 Biochemical Society
Imaging plant virus infections
may influence localization, for instance a protein’s ability to move
through PD (plasmodesmata) [57] or nuclear pores [58]. Some
FP-fusions are unstable, and proteolytic processing may release a
significant pool of the free FP resulting in nucleo-cytoplasmic
labelling [7,39,59]. When this is suspected, Western blotting
against GFP should be carried out to test for degradation of the
fusion protein. If mistargeting is suspected, valuable information
may still be gleaned from the data. For instance, a secondary or
transient localization may only become apparent when the main
targeting site of a protein of interest is disrupted. This is especially
relevant in the case of multifunctional viral proteins. In most cases,
the orientation of the FP-fusion protein will have the greatest effect
on localization. In a recent study of the TGB MPs of grapevine
rupestris stem pitting-associated foveavirus, Rebelo et al. [60]
systematically investigated the effects of FP-fusions to each TGB
protein on both N- and C-termini. Such a thorough approach
may not always be necessary, but with the available repertory of
Gateway fusion vectors it can be done relatively quickly. If only
one fusion orientation is used, publications should clearly state
which, and for what reason it was selected. On some occasions, the
best approach may be to introduce the fluorescent tag internally at
a boundary between different protein domains [61]. Few reports
using such strategies have been published in plant virology to date
(e.g. [62]), probably because there is surprisingly little structural
information available for most plant viral proteins. An approach
for rapid production of internal FP-fusions based on the Gateway
technology has been described [61].
Some of these difficulties might be overcome if smaller, less disruptive fluorescent tags were available, which would additionally
help to limit the detrimental effects of introducing the tag into viral
genomes. One novel FP that meets some of these requirements
is iLOV [63], derived from the light-, oxygen- or voltagesensing LOV2 domain of the Arabidopsis thaliana photoreceptor
phototropin 2. With a size of ∼ 300 bp and 17 kDa, iLOV is
approx. 1.5–2-fold smaller than GFP and has been demonstrated
to outperform GFP in some virus-specific contexts [63]. For
example, TMV (tobacco mosaic virus) expressing unfused or
MP-fused iLOV moved faster cell-to-cell and systemically than
the corresponding GFP constructs, indicating advantages both in
terms of genetic load and functionality of the MP-fusion.
If a protein-specific antibody is available, immunofluorescence
microscopy can make tagging altogether unnecessary, but it is
limited to fixed samples. A potential compromise permitting livecell imaging could be to use a small non-fluorescent tag that is
rendered fluorescent only by addition of a specific interacting
compound (probe). A variety of such tag–probe labelling
systems have been developed (reviewed in [64–66]), but
unfortunately the majority are cell-impermeant and therefore
unsuitable for plant studies. Of those which permit intracellular
labelling, the proprietary SNAPTM and CLIPTM tag systems based
on O6 -alkylguanin-DNA alkyltransferase (New England Biolabs;
[67,68]), HaloTagTM (Promega; [69]), based on haloalkane
dehalogenase, and LigandLinkTM , based on Escherichia coli
dihydrofolate reductase (ActiveMotif; [70]), all utilize the
reaction between small enzymes and substrate analogues to link
a fluorophore covalently to a protein of interest, but provide only
a modest size advantage over GFP (27 kDa) with tag molecular
masses between 18 and 33 kDa. Only HaloTagTM has been tested
in plants, with good tissue permeability [71]. A tag based on
the human immunophilin FKBP12 (FK506-binding protein 12)
[72,73] adds only 12 kDa, but has not been tested in plants
yet. The smallest available tags with cell-permeant fluorescent
probes are a 38-amino-acid peptide that binds Texas Red with
picomolar affinity [74], which also has not yet been tested
in plants, and the tetracysteine tag/biarsenical ligand system
25
[75,76]. The optimized, tetracysteine-containing peptide tag is
of similar size as an epitope tag (12 amino acids [77]). The
biarsenical probes FlAsH (fluorescein arsenical helix binder) and
ReAsH (resorufin arsenical helix binder; [78]) are membranepermeant non-fluorescent fluorescein derivatives that become
fluorescent only when covalently bound to the tetracysteine tag.
This ‘switch on’ is an advantage over the Texas Red-binding
tag, and particularly useful as it eliminates the washing out of
unbound dye, which is time-consuming and a problem when
the tissue needs to be studied rapidly. Despite its promise, the
tetracysteine tag/FlAsH system has seen little use in plants so far
[79]. The biarsenical probes can cause cytotoxic effects [76] and
some background staining may result from non-specific reactions
with other cysteine-containing proteins [64,80]. From our own
experience, the greatest problem in plants may be insufficient
permeability through (and non-specific labelling of) the cell wall
(P. Boevink, N.M. Christensen and K.J. Oparka, unpublished
work). Tag–probe labelling systems could be an ideal approach to
localize fusion-sensitive viral proteins, but so far none is entirely
suited to plant systems.
TRACKING PROTEIN DYNAMICS AND LOCALIZING TRANSLATION
SITES
Many proteins are not static in the cell and some may
take different pathways to the same subcellular localization.
Therefore significant functional information can be gained from
analysing protein dynamics. With standard FP-based fluorescence
microscopy, this is possible only to a limited extent, e.g. by
following protein localization over a time course. A range of
techniques has been developed that permit more detailed insights
into protein transport pathways, mobility and also the site of their
synthesis. Tracking approaches are particularly valuable in the
study of multifunctional viral proteins during an infection cycle,
while determining the site of viral protein translation may help to
link viral replication and protein function.
FRAP (fluorescence recovery after photobleaching) and FLIP
(fluorescence loss in photobleaching)
One way to gain dynamic information from FP-fusion localization
studies is to use photobleaching approaches [81,82]. Highexcitation light intensities ‘overload’ the energy absorbtion of
fluorescent molecules and destroy them permanently, leading
to photobleaching. With the spatially restricted light intensities
that can be achieved with lasers, it is possible to locally
bleach a specific fluorophore within a cell in a selected ROI
(region of interest). Because all cellular proteins, including
FP-fusions, participate in dynamic interactions with their
environment, the non-fluorescent bleached molecules from the
ROI will subsequently mix with their still-fluorescent counterparts
elsewhere in the cell. The kinetics of this process can be measured
and used to draw conclusions about the mobility and transport
pathways of FP-fusion proteins. In FRAP experiments, the return
of fluorescence to the bleached area is monitored. If the FPfusion protein is freely diffusive within the cell, fluorescence will
return rapidly and with a simple kinetic based on cellular diffusion
constants. If the protein has limited mobility, e.g. by interactions
with other cellular macromolecules, fluorescence recovery will be
slowed and will follow a more complicated kinetic. For instance,
Goodin et al. [49] were able to distinguish two intranuclear
pools of the N (nucleocapsid) protein of Sonchus yellow net
rhabdovirus. One pool of N in the nucleoplasm showed fast FRAP
kinetics and was probably present in a soluble form. The other pool
c The Authors Journal compilation c 2010 Biochemical Society
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J. Tilsner and K. J. Oparka
was membrane-associated and showed very slow FRAP kinetics,
indicating that the protein is immobilized. N interacts with another
viral protein, P, but FRAP experiments on N/P-co-expressing
uninfected cells showed similar kinetics to soluble N, indicating
that viral factors other than P are responsible for immobilizing
N on membranes. If the bleached pool of an FP-fusion is
compartmentally isolated, no fluorescence recovery may occur
at all. Goodin et al. [49] used this FRAP approach to show that
the membranous intranuclear replication sites of Sonchus yellow
net virus are continuous with the nuclear envelope. Both the
replication sites and the inner nuclear envelope were labelled by
a GFP-fusion of the human lamin B receptor, and photobleached
replication sites quickly recovered their fluorescence, indicating
redistribution from the connected cellular endomembranes.
FRAP can also give information on the trafficking pathway a
FP takes to reach the bleached area. Wright et al. [83] combined
FRAP of the PD-localized TMV 30K MP with cellular inhibitors
to analyse the intracellular route of the protein to PD. Microtubuledepolymerizing drugs had no effect on 30K PD targeting to
PD, whereas actin cytoskeleton inhibitors significantly inhibited
MP targeting (Figure 1c). Also, high concentrations of brefeldin
A sufficient to distort the cortical tubular ER network and
treatment with the general metabolic inhibitor sodium azide both
significantly reduced fluorescence recovery at PD, indicating that
the 30K protein is actively transported along the cortical ER/actin
network.
In contrast with FRAP, the depletion of fluorescence from
the surrounding non-bleached cell areas is monitored in FLIP
experiments. Similar to FRAP, this allows conclusions to be
drawn about protein-trafficking pathways as well as connectivity
between compartments.
Fluorescent highlighter proteins and tag–probe pulse–chase
labelling
FRAP and FLIP are not suitable for continuous tracking of the
same protein pool or for distinguishing newly synthesized from
pre-existing protein. The latter is of particular interest to the study
of virus infections because the location of viral protein synthesis
is of considerable functional relevance. The native location of
viral proteins in the course of infection may result from both their
intrinsic properties and their site of synthesis. For instance, plant
virus MPs must associate with the genome they are transporting,
but so far none has been characterized as sequence-specific nucleic
acid binders [4]. In some cases, specific interactions between the
MP and other viral proteins, such as CPs, may aid specificity
(e.g. [84]), but the spatial proximity of the MPs to progeny
genomes, i.e. ‘co-compartmentation’, may also play a role which
could be achieved by synthesis and retention of the MP in the
vicinity of the replication site. Clear identification of a specific
subpopulation of proteins in a cell requires differential labelling.
This can be achieved using either FPs whose spectral properties
can be changed in vivo, so-called photoconvertible or ‘highlighter’
FPs (pc-FPs; reviewed in [18,64,85,86]). Alternatively, tag–probe
systems can be employed that permit pulse–chase labelling with
differently coloured probes. Both approaches permit the selective
highlighting of the most recently synthesized protein pool and can
thus identify translation sites. The ability to change the colour of
the fluorescently labelled protein locally in a subcellular ROI
and track its movement through the cell is, however, limited to
pc-FPs.
One group of pc-FPs, including PA (photoactivatable)-GFP and
PA-mCherry, is switched irreversibly from a near-non-fluorescent
dark state to green or red fluorescence respectively by strong UV
c The Authors Journal compilation c 2010 Biochemical Society
illumination [18,64,85,86]. Activatable pc-FPs permit selective
tracking of subcellular molecule populations, but do not allow
pulse–chase labelling. Additionally, they are difficult to detect
prior to photoactivation, which therefore has to be done ‘blindly’
or using other cellular markers for orientation. A different group
of pc-FPs are ‘switched’ on and off reversibly, and include the
variants of Dronpa, Padron, KPF1 (Kindling), rsFastlime and
rsCherry, IrisFP and DsRed Timer (see reviews [18,64,85,86]
for comparisons of spectral properties). These proteins permit
repeated on/off switching (in some cases many cycles), and can be
used for repetitive FRAP or tracking experiments within the same
cell. They are also important for new subdiffraction-resolution
microscopy approaches (discussed below).
The most relevant group of pc-FPs for pulse–chase labelling are
true photoconverters, i.e. irreversibly colour-changing proteins.
These include PS-CFP2 (where CFP is cyan FP), which switches
from cyan to green fluorescence, and numerous green-to-red
switching pc-FPs: Kaede, KikGR (Kikume), EosFP variants
and Dendra2; also KFP1 (kindling FP1) can be green-intored converted irreversibly (for details of spectral properties, see
[18,64,85,86]). Additionally, it was recently discovered that many
red FPs can be photoswitched, including Katushka, mKate and
HcRed1 (from red to green) and mOrange1 and 2 (from red to farred), making them suitable as highlighters [87]. A conversion into
cyan fluorescence has also been described for the YFPs (yellow
fluorescent proteins) EYFP (enhanced YFP), Venus and Citrine
[87,88].
Colour-changing pc-FPs are ideally suited for both tracking
and pulse–chase labelling experiments. The label is easily visible
before photoconversion, and the colour change leaves both
switched and unswitched protein pools visible simultaneously.
Movement of selected particles, organelles etc. can thus be
followed, whereas newly synthesized protein can be visualized
by first (irreversibly) photoswitching the entire pc-FP population
in a cell and then observing reappearance of the unswitched colour
due to de novo translation. This approach has been used to identify
localized translation sites of a potassium channel in neuronal
dendritic appendages [89] (Figure 1d). Translation inhibitors such
as cycloheximide and puromycin can be used for controls. Also,
both FRAP and PA-GFP were used to show that the NS4B and
NS5A proteins of hepatitis C virus remain associated with putative
viral replication sites when expressed from the virus, but are
motile in uninfected cells [90,91]. One potential problem with the
use of pc-FPs in plants may be that the experimental organism
has to be kept in the light. Since natural daylight contains a
UV component, this can lead to undesired photoswitching (some
pc-FPs are also switched by visible wavelengths). A potential
solution is to keep the plants in the dark for some time before
microscopy.
Pre-existing and newly synthesized protein pools can also be
pulse–chased using tag–probe labelling systems. The existing
protein pool is first labelled with one fluorophore, then subsequent
labelling with a different fluorophore selectively highlights
only the most recently synthesized protein, identifying the
translation sites. The tetracysteine tag/biarsenical ligand system
has been used to localize β-actin translation sites in vivo with
FlAsH/ReAsH pulse–chase labelling [92]. Among the other tags
which can be used intracellularly [64–66], the HaloTagTM [69],
SNAP/CLIP-tagTM [67,68] and LigandLinkTM [70] systems also
offer different probe colours for pulse–chase labelling. However,
a disadvantage over FlAsH/ReAsH is that these probes are
permanently fluorescent. Non-fluorescent probes are available
for blocking the pre-existing protein pool instead of changing
the label colour, but the tissue still has to be washed to remove
non-specific fluorescence. Lang et al. [71] have reported washing
Imaging plant virus infections
times of between 4 and 18h for HaloTagTM , the only one of
these systems tested in plants. Unless the tissue is fixed, these
times are too long for detecting translation sites. Lin et al. [93]
recently introduced another alternative technique, TimeSTAMP,
for selective labelling of translation sites. The protein of interest
is tagged with an epitope tag via a linker that encodes both the
NS3 protease from hepatitis C virus and its NS4A/B cleavage
site. Autoproteolysis constantly removes the tag until a specific
membrane-permeant protease inhibitor is added. Any protein
synthesized after addition of the inhibitor will remain labelled
and can be localized by immunofluorescence microscopy. The
permeability of the inhibitor through plant tissue remains to be
tested.
DETECTION OF PROTEIN–PROTEIN INTERACTIONS
Subcellular localization of a protein needs to be put into a spatial
and functional context to permit conclusions about its role during
infection. Co-localizations with other proteins including organelle
markers can provide such a context. However, co-localizations do
not provide proof of macromolecular interactions. The resolution
limit of confocal microscopes in the focal plane is approx. 200–
250 nm and even lower in the z-axis. A spherical volume with a
diameter of 200 nm may contain up to 140 000 GFP molecules
that could be co-localized without any physical interaction [94].
Only in special situations can co-localization be a sufficient
indicator of protein interactions, for instance when one FP-fusion
causes a clear redistribution of another, or when this is coupled
to a functional phenotype. Therefore fluorescence microscopy
applications have been developed to more specifically identify
protein–protein interactions in vivo.
BiFC (bimolecular fluorescence complementation)
BiFC, reviewed in [39,94–96], is the reconstitution of a FP
from two non-fluorescent fragments. BiFC can be used to detect
protein–protein interactions because FP reconstitution depends
on the FP halves being in close proximity long enough for
refolding, a process thought to require up to several minutes. To
detect interactions, the split-FP halves are translationally fused
to two proteins of interest, whose interaction then facilitates
BiFC, thereby ‘switching on’ fluorescence. In contrast with other
protein fragment complementation approaches such as the yeasttwo-hybrid system, BiFC not only allows direct microscopic
visualization of the interaction but also provides additional
information on the localization of the interaction within the
cell. As for conventional FP-fusions, several sets of plant BiFC
vectors compatible with restriction cloning [97–101] or Gateway
technology [30,31,98,102] have been developed to facilitate rapid
and easy experiments.
FPs are commonly split at either of two suitable splitting
positions, between amino acids 154 and 155 or 172 and 173
respectively (actual positions may vary slightly depending on the
FP variant) [95]. In both cases, the N-terminal fragment (FPN)
is larger and contains the side chains forming the fluorophore.
The smaller FPC (C-terminal fragment of split FP) is probably
required to close the β-barrel tertiary structure that protects the
fluorophore and allows its maturation. There do not seem to
be any obvious benefits or disadvantages between the 154/155
and 172/173 splitting positions and they are being used rather
indiscriminately at this point. However, FPN172 fragments can
be complemented by FPC154 counterparts, whereas FPN154
fragments cannot be complemented by FPN173 halves [103],
indicating that the refolded FP structure tolerates a small peptide
‘overlap’, but not a gap. This is of practical relevance as cross-
27
complementation between different split-FP variants leads to
the formation of FPs with intermediate spectral properties and
thus allows simultaneous detection of different protein–protein
interactions in the same cell [103]. Multicolour BiFC can, for
instance, be used to test two putative or competing interaction
partners (fused to different FPN fragments) of the same bait
protein (fused to an FPC). Cross-complementation competencies
and resulting spectral properties of the hybrid FPs are reviewed in
[95,103]. Notably, GFP-derived FPC does not complement any of
the other FP variants tested. Multicolour-BiFC has been used in
plants, and suitable vector sets are available for both restrictionbased [100,101] and Gateway [31,102] cloning, so this developing
technique awaits exploitation by plant virologists.
Because of its relatively straighforward use, BiFC has quickly
become the most popular technique to detect protein–protein
interactions in vivo, and has been used in numerous plant
virus studies. It is therefore worth being aware of the method’s
complications and pitfalls, which are also reviewed in [94] and
[95]. A general drawback of BiFC is that the FP reconstitution
is practically irreversible, making the technique unsuitable for
monitoring dynamic interactions. This irreversibility potentially
increases BiFC sensitivity, although it can also result in the
accumulation of non-specific background fluorescence. For
several experimental systems, no fluorescence was reconstituted
when unfused FPN and FPC fragments were co-overexpressed.
However, for some FPs, background fluorescence has been
observed (e.g. [62,94]). Li et al. [104] have recently found that
shortening the N-terminal fragment of YFP split at position 154 to
YFP-N152 eliminates non-specific background, albeit at the cost of
reduced signal intensity. In any case, unfused split-FP fragments
are not a sufficient control in BiFC experiments, as the fusion to
proteins of interest may influence their folding and stability so that
non-specific fluorescence occurs for fused FP fragments even if
no background was observed with unfused FPN+FPC [94,105].
Therefore fusions to known non-interacting proteins are a more
adequate control. Additionally, the splitting position and the
orientation of the split-FP-fusions may influence both undesired
background signal and the detection of genuine interactions.
The sterical constraints imposed on the FP reconstitution by
fused interacting partners appears to be complex (see [94]).
Sometimes one orientation will yield fluorescence, whereas
another will not [62,97], and sometimes fusion orientations have
no effect. Including information on splitting positions and fusion
orientations in publications may help to improve the comparability
between different experiments. Finally, both non-specific FP
complementation and relative brightness of the BiFC signal will
be increased by molecular crowding, i.e. if the FP-fusions are
confined to a small subcellular compartment. Studies of such
compartments require particularly rigorous controls, and the
use of strong promoters is best avoided in BiFC experiments.
However, even non-specific background fluorescence in BiFC
can be used to obtain valuable information about the topology of
integral membrane proteins (see below).
FRET (fluorescence resonance energy transfer)
FRET is the direct, nonradiative transmission of excitation energy
from one fluorophore (FRET donor) to another (FRET acceptor),
resulting in fluorescence of the acceptor after excitation of the
donor. For this, the fluorophores have to come into very close
proximity (10 nm), as FRET efficiency decreases with the sixth
power of distance. This makes it ideally suited to study molecular
interactions (reviewed in [39,94,106]). For this purpose, similar to
BiFC, the FRET partners are linked to the macromolecules whose
interaction is to be studied. In contrast with BiFC, which is based
c The Authors Journal compilation c 2010 Biochemical Society
28
J. Tilsner and K. J. Oparka
on the properties of a specific protein tertiary structure, FRET
can occur between any kind of fluorophore, and is therefore much
more versatile. For instance, it can be used to study interactions
between proteins and other molecules such as nucleic acids
[107]. FRET can also occur intramolecularly between different
fluorophores attached to the same macromolecule, allowing
microscopic study of conformation changes (e.g. [108]). Perhaps
most importantly for in vivo applications, the interaction between
the FRET fluorophores themselves remains transient so that no
non-specific background signal can accumulate. However, nonspecific FRET can result from random collisions of fluorophores,
and this probability increases with fluorophore concentrations and
decreasing experimental volume. FRET measurements therefore
always require adequate controls.
In the most straightforward method for FRET detection,
the donor is excited with an appropriate wavelength, and
fluorescence emission is then detected in the spectral window
of the acceptor (‘sensitized emission’). For this approach,
well-separated excitation and emission spectra respectively of
donor and acceptor are desirable to avoid bleedthrough (donor
fluorescence in the acceptor channel or direct excitation of the
acceptor by donor-‘specific’ wavelengths). On the other hand,
efficient FRET requires sufficient overlap between the donor
emission and acceptor excitation spectra. Controls that are good
practice for FRET experiments of this type include measurement
of FRET efficiency for the non-interacting (unfused) FRET pair
(background level due to random collision), fluorescence intensity
measurements of the individually expressed FP-fusions with both
donor- and acceptor-specific filters (to quantify bleedthrough) and
ideally also a positive control, usually a direct fusion between
donor and acceptor FPs. An alternative approach, which can
also be used as an additional control in sensitized emission
experiments, is acceptor photobleaching. Here, the quenching
(reduction) of donor fluorescence by interaction with the FRET
acceptor is measured. Selective photobleaching of the acceptor
restores the full fluorescence intensity of the FRET donor
(unquenching), allowing quantification of FRET efficiency. For
this, the spectral properties of the FRET pair need to permit
selective bleaching of only the acceptor. All of these methods
depend on the measurement of light intensities, i.e. photon counts,
and are therefore dependent on fluorophore concentrations.
However, reproducible, consistent levels of the same construct are
difficult to achieve in independent in vivo experiments, leading
to a certain level of inaccuracy in all intensity-based FRET
measurements.
The most accurate method for FRET detection is FLIM
(fluorescence lifetime imaging) of the FRET donor [109,110].
Energy transfer to the FRET acceptor shortens the average
time that a donor molecule remains in the excited state, which
for FPs is usually in the order of nanoseconds. Reduction
of donor fluorescence lifetime is independent of fluorophore
concentrations and depends only on the FRET pair characteristics
and fluorophore distance. FLIM requires sophisticated special
equipment: for ‘time-domain’ FLIM, where donor fluorescence
decay is measured directly, a pulsed laser capable of producing
bursts of excitation light sufficiently shorter than the decay times is
required, as well as similarly fast time-resolved photon detection.
‘Frequency domain’ FLIM measurements use an excitation laser
that produces continuous light with a sinuosidally modulating
intensity. Fluorescence lifetime is deduced from the phase-shift
between the intensity curves of excitation and emission fluorescence, again requiring specialized detection equipment. Because
of these technical demands and prohibitive costs, FLIM–FRET,
although considered the ‘gold standard’ for microscopic detection
of protein–protein interactions, is not yet widely available.
c The Authors Journal compilation c 2010 Biochemical Society
FRET experiments are also dependent on the choice of
FRET pair. Many FP pairs have been used for FRET, and a
complete listing is beyond the scope of the present review (see
[17,18,39,94,106,111] instead). The most commonly used FRET
pair, CFP/YFP, has a number of problems. YFP has a small
Stokes shift (distance between excitation and emission maxima),
making it harder to avoid CFP bleedthrough. Additionally,
EYFP can change into a CFP-like blue form in response to
intense UV light during acceptor photobleaching, leading to
overestimation of the FRET contribution to CFP unquenching
[88]. This effect has been disputed [112], but was recently also
reported for the YFP variants Venus and Citrine [87]. CFP is
comparatively dim and its fluorescence decay curve follows
a double exponential, complicating FLIM quantification [113].
Brighter CFPs with single-exponential fluorescence decay are
available [94,114]. Recently, TagGFP/TagRFP (where RFP is
red FP) has been promoted as an optimized FRET pair [115].
Importantly, in contrast with several other RFPs, TagRFP shows
no photoconversion under strong UV illumination, avoiding
potential artefacts during acceptor photobleaching [87]. Also
of note is a non-fluorescent YFP variant, REACh [116], which
permits FLIM- and intensity-based FRET measurements with a
GFP donor, free of any spectral bleedthrough.
Perhaps due to the technical sophistication required for FRET
experiments, BiFC has so far proven the more popular method
to study protein interactions in vivo among plant virologists.
Recently, both approaches have been combined to study ternary
protein interactions in planta [117]. However, because of its
reliance on transient molecular interactions and applicability
to any type of fluorophore, FRET has tremendous potential
for the functional study of plant virus replication, including
protein–protein interactions within the replication site (e.g.
[118]), protein–RNA interactions ([107,119]) and potentially even
RNA secondary structure changes (discussed below). Ideally, both
BiFC and FRET protein–protein interaction experiments should
be verified by genetic approaches such as yeast two-hybrid or
split-ubiquitin complementation assays.
ANALYSIS OF PROTEIN MEMBRANE TOPOLOGY
Many plant viruses encode integral membrane proteins, including
various MPs [4]. So far, very few molecular structure data are
available for these proteins. Of particular interest is information
on the topology of viral transmembrane proteins, to gain
insights into the position of functionally important groups
with regard to the cellular surroundings and thus potential
interaction partners. Integral membrane proteins may have one
or several hydrophobic transmembrane helices and, depending
on their mode of membrane insertion, the N-terminus can be
directed towards the cytoplasm or towards the lumen of the ER
(corresponding to the apoplast for plasma membrane proteins),
also determining the orientation of other transmembrane domains
and connecting loop regions. Membrane topology can be
analysed by immunofluorescence microscopy. A tag is fused
to either end of the transmembrane protein (or internally) and
its accessibility to antibody labelling at the cell surface (nonpermeabilized) and intracellularly (membrane-permeabilized)
is compared (e.g. [120]). However, this is unsuitable for
intracellular endomembrane-resident proteins. Additionally, it
requires fixation and permeabilization. Analogous biochemical
approaches, e.g. using protease accessibility, require disruption
of the cell. None of these approaches permits analyses in vivo.
Two new live-cell imaging methods have been developed for
analysis of membrane topology. Zamyatnin Jr et al. [62] used the
Imaging plant virus infections
non-specific fluorescence that occurs in BiFC assays: if one half of
a split FP is fused to a transmembrane protein, the complementing
split FP fragment then has to be localized to the same compartment
for restoration of fluorescence. They demonstrated that a FPN
fragment fused to either of the N- or C-termini of the potato moptop virus TGB2 protein, which has two transmembrane domains,
reconstituted fluorescence only in combination with cytosolic
FPC. On the other hand, an internal fusion of FPC within the loop
region connecting the transmembrane helices led to BiFC only
in combination with FPN targeted to the ER lumen. Accordingly,
both N- and C-termini are located on the cytosolic face of the ER
and the loop domain in the lumen (Figure 1f). This approach
could also be modified for plasma membrane transmembrane
proteins: only their cytoplasmic domains will enable BiFC with
intracellular split FP fragments. The authors did find, however,
that unfused cytoplasmic FPN was able to trap the smaller
ER-targeted FPC in the cytoplasm, resulting in cytoplasmic
fluorescence. Therefore they used only the larger ER-targeted
N-terminal YFP fragment to detect ER-luminal BiFC [62].
An alternative method uses the redox-sensitivity of a GFP
variant together with redox differences in the cytoplasmic and ERluminal compartments, and is therefore limited to endomembraneresident transmembrane proteins (ReTA, redox-based topology
analysis; [121]). In this approach, the transmembrane protein
of interest is fused to the redox-sensitive GFP variant roGFP2
[122], which is strongly excited at 405 nm in its oxidized form,
but only very weakly excited in the reduced state. The ratio of
fluorescence after 405 nm and 488 nm excitation can thus be
used to determine the position of roGFP2 in the cytoplasm (more
reducing, low 405/488 nm excitation ratio) or the ER lumen (more
oxidizing, high 405/488 nm excitation ratio). When fused to a
membrane protein of interest, the 405/488 nm excitation ratio
reveals the orientation of the fused protein domain in the ER or
cytosol (Figure 1g). These new tools, together with the analysis
of protein–protein interactions in vivo will greatly facilitate our
understanding of the membrane-associated protein complexes
involved in plant virus infections.
RNA IMAGING
Since the advent of molecular cloning in the 1970s, the functional
investigation of viral genomes, including sequence determination,
analysis of expression strategies and identification of regulatory
sequence elements [42,43], has been at the forefront of modern
virology. The link between such data and the current wave of
FP studies is the location of the viral genome in relation to its
encoded proteins and host cell structures. Where, and in what
cellular context, do the genetically characterized infection events
actually take place? In situ hybridization at the light and electron
microscopic level can provide some of this information, but is
experimentally demanding and reveals only snapshots of dynamic
processes. RNA localizations by in situ hybridization have often
been carried out on protoplast systems due to better permeability
and because some temporal resolution can be achieved by
stopping synchronized infected cultures at defined timepoints
(e.g. [123,124]). However, all RNA viruses replicate on cellular
membranes which they reorganize extensively into a VRC (viral
replication centre) [3,5] and the sensitivity of cellular membranes
to standard aldehyde fixation protocols commonly employed in
in situ hybridization is another reason for making in vivo RNA
tracking a preferred choice.
Tools for sequence-specific live-cell RNA detection have been
available since 1998 (reviewed in [125,126]). The first such
system, based on the coat protein of bacteriophage MS2 [127],
and a more recent version using a 22-amino-acid peptide from
29
the N protein of bacteriophage λ (λN22 ) [128], employ fusion of
a sequence-specific RNA-binding protein to GFP. The MS2 CP
and λN22 peptides recognize RNA hairpins with which the RNA
of interest needs to be tagged. In order to separate unbound and
RNA-bound reporter, the GFP-fusion is usually targeted to the
nucleus. Cytoplasmic fluorescence then indicates redistribution
in response to RNA binding. The MS2 system has been used
in numerous studies, and MS2 and λN22 together have enabled
simultaneous tracking of two RNAs in the same cell [129]. Zhang
and Simon [130] tracked turnip crinkle virus infection using the
MS2 system, but did not localize the viral RNA at the subcellular
level, although the technique is principally suited to this. Sambade
et al. [131] showed that a nuclear expressed, MS2-tagged RNA
encoding the TMV MP formed motile granules in tobacco leaf
epidermal cells which co-localized with MP at PD. A limitation
of the MS2 and λN22 systems for virus imaging is the introduction of additional sequences with extensive secondary structure
into the viral RNA, which may affect infectivity, recombination
rates or RNA localization [43,126,129]. Additionally, multiple
tandem copies of the hairpin tag have to be introduced to obtain
sufficient sensitivity (commonly 6–24, but up to 96 repeats have
been used; [125]). Zhang and Simon [130] used only a single
MS2 hairpin to tag turnip crinkle virus, which was sufficient for
relocalization of MS2 CP–GFP from the nucleus to the cytoplasm,
but this may permit subcellular RNA localization for very highly
accumulating viruses.
Because of these limitations, a novel approach based on
the RNA-binding domain of the translational repressor human
Pumilio1 [132] held promise for virus studies. The PUMHD
(Pumilio homology domain) has a modular structure of eight
imperfect repeats that each bind one nucleotide with just three
amino acids per repeat involved in the protein–RNA interaction.
This makes it possible to alter the sequence specificity of the
PUMHD in a defined way with relatively little genetic engineering
of the protein [133,134]. Instead of tagging the viral RNA, the
detection system can be modified to recognize the unaltered RNA.
Additionally, the PUMHD has a higher affinity (K d = 0.48 nM)
to its target sequence than either MS2 CP (6.2 nM) or λN22
(22 nM; reviewed in [126]), and binding neither requires nor
introduces RNA secondary structure. Also, the need for nuclear
sequestration of the unbound reporter was avoided by instead
using BiFC between two PUMHD variants engineered to bind
to closely adjacent target sites on the same RNA and fused to
the two halves of a split FP [132]. This system enabled detailed
mapping of the distribution of unencapsidated RNA of TMV and
PVX (potato virus X) in their respective VRCs (Figure 1h), as
well as the co-introduction of an RNA reporter and a 35S-driven
TMV replicon to study early vRNA (viral RNA) distribution
beginning ∼ 6 h after co-bombardment [105]. The perinuclear
VRC of TMV contained granular RNA ‘hot spots’, whereas
that of PVX showed circular RNA ‘whorls’ arranged around
aggregates of the viral MP TGB1, indicating that these structures
show considerably different arrangements in different viruses.
The PUMHD variants were modified to recognize unaltered TMV
genomic RNA, but subsequent tagging of the PVX genome with
the same TMV-derived sequences demonstrated that this short
(21 nt) non-structured tag had no disruptive effect on the virus
[105]. The PUMHD-BiFC reporter has also been used to localize
potyviral RNA to replication sites in membrane invaginations of
the chloroplast envelope [36].
More recent structural studies of the PUMHD protein have
revealed that its sequence specificity is not as high as initially
believed [134]. A certain degree of binding promiscuity is
probably balanced by the requirement of BiFC on binding of two
PUMHD fusions to generate a signal, but the BiFC approach
c The Authors Journal compilation c 2010 Biochemical Society
30
J. Tilsner and K. J. Oparka
is itself not free of background signal. Additionally, altering
PUMHD specificity often results in a marked reduction in affinity
[132,133] and thus sensitivity. PUMHD-BiFC has enabled the
most precise in vivo localizations of viral RNA genomes so far,
but further developments are probably required to provide plant
virologists with a generally applicable live-cell RNA reporter.
Extremely high sensitivity and signal-to-noise ratios can be
achieved by methods that require invasive delivery of the RNA
reporter into cells. In direct RNA labelling, the viral genome
itself is rendered fluorescent by incorporation of fluorophorecoupled nucleotides during in vitro transcription and then
microinjected [135]. Although low-throughput, this approach
overcomes the temporal limitations inherent in virus-expressed
FP reporters, and permits insights into the very early stages
of infection. Microinjected TMV RNA was found to attach
to the ER/actin network immediately after entry in a capdependent manner [135], possibly in connection with the host
translation machinery [136]. Microinjection of directly labelled
viral RNA encapsidated in fluorescence-conjugated CP allows the
visualization of virus unencapsidation in vivo, also revealing the
cellular context in which this infection step happens [135,137].
It is conceivable that such approaches could be expanded
further, utilizing FRET to study changing macromolecular
interactions during early infection events [119]. For instance,
FRET between an RNA-intercalated dye and a protein FPfusion was recently used to detect protein–RNA interactions
in vivo [107]. Similarly, directly labelled viral RNA could
be microinjected into cells expressing an FP-fused replication
factor to study early interaction events. Alternatively, molecular
beacons could be used. Molecular beacons are short synthetic
oligonucleotides linked to fluorophores that are used for sequencespecific RNA detection in vivo and in vitro (reviewed in
[125,126,138]). Molecular beacon signal is limited to the targetbound form because in the unbound state secondary-structure
formation of the beacon brings a fluorophore and a quencher
group in close contact, preventing fluorescence emission [139].
The beacons could be microinjected to detect viral RNAs, but
another approch would be to hybridize them to the viral genome
in vitro and then microinject beacon-labelled vRNA. The binding
of the molecular beacon is sensitive to secondary structure of the
target RNA [125,126,138,140]. Therefore disapperance of
the fluorescent signal could be used to monitor displacement of the
molecular beacon by RNA secondary-structure rearrangements or
protein binding in vivo. Protoplasts would probably be the easiest
system for such experiments.
For some questions, no in vivo imaging techniques
are currently available. For instance, double-stranded RNA
replication intermediates are a particularly good indicator of
replication sites. Currently they can only be visualized by
immunofluorescence microscopy with dsRNA (double-stranded
RNA)-specific antibodies (e.g. [123,124]), although the use of
preferentially dsRNA-binding cell-permeant dyes may provide
an in vivo alternative [137]. Newly synthesized RNA can
be visualized by incorporation of bromo-UTP and subsequent
detection with specific antibodies (e.g. [123,124]), an approach
that is also suitable for DNA viruses, in which bromo-dUTP
is used instead (e.g. [53]). With the available toolbox of RNAimaging techniques, genetic and microscopical plant virus studies
can now be linked at the subcellular level.
TRACKING THE COMPLETE VIRAL LIFE CYCLE IN FOUR
DIMENSIONS
For some animal viruses, the complete viral lifecycle can now be
followed microscopically (e.g. [52,141]). All viral gene products
c The Authors Journal compilation c 2010 Biochemical Society
can be localized throughout infection either in living cells or cell
cultures fixed at specific time points. Viral nucleic acids can be
visualized and the interactions of viral components with host cell
factors and among each other can be detected in vivo, individual
viral genomes or particles can be tracked [142], and even transient
and complex events such as virus entry [137,143], packaging
[144] and exit/cell–cell-transfer [145] can be observed. For plant
systems, the same tools and methods are also available, with
certain imaging limitations due to autofluorescence from the cell
wall and chloroplasts [64]. It can be expected that the spatial and
temporal relationships between plant viruses and their host cells
will soon be understood in similar detail.
The earliest events in the viral infection cycle are the most
difficult to visualize. Usually, only a few viral genomes and
a very low level of their encoded proteins are present in the
host cell. Additionally, virus-expressed fluorescent reporters
only become detectable after a few hours, due to a lag in
expression and fluorophore maturation. One way to study early
events, such as unencapsidation of virions and the initiation
of translation, is to label viral components in vitro and then
introduce them invasively, which may not differ all that much
from the ‘natural’ delivery of plant viruses via mechanic wounds
or insect feeding tubes. If unincorporated fluorescent molecules
are removed after labelling, in vitro labelling yields extremely
high signal/noise ratios. Also, larger pools of virions can be
introduced than would be the case in a natural infection. This
may lead to artefacts in some cases, but also overcomes some
of the limitations resulting from low virus levels. Studies such
as those of Christensen et al. [135], which investigated the
initial localization and unencapsidation of microinjected CP- and
RNA-labelled TMV virions could be expanded to RNA–protein
and protein–protein interaction dynamics by FRET, as discussed
above. Alternatively, transfection of entire double-labelled virus
particles into protoplasts might be possible using electroporation,
poly(ethylene glycol) precipitation or membrane-permeabilizing
agents.
Very early infections events can also be studied indirectly.
Cotton et al. [146] infected the same cells with two different
potyviral genomes expressing either GFP- or RFP-fusions to
the 6K protein that establishes the replication complex on the
ER. The replication complexes were either green or red, rarely
both, leading the authors to conclude that each replication site
is established from a single infectious genome and that the p6
protein remains associated with the same replication site. Not
all viral systems permit double infections, but where this is
possible, 35S promoter-driven infectious clones are particularly
suitable as different viruses can be co-introduced into the same cell
with nearly 100 % co-transformation efficiency through particle
bombardment [147].
Large imaging gaps are also still to fill in two areas where
plant and animal viruses show the greatest discrepancies. These
are virus entry and exit, both between neighbouring plant cells
and between the host plant and vector organisms. Entry into
and exit from the plant are usually achieved mechanically, i.e.
through perforation of the cell wall, whereas cell-to-cell transport
within the host organism occurs through the plasmodesmal pores
traversing the walls. By contrast, the majority of animal viruses
are membrane-coated and enter and leave cells by membrane
fusion and budding events. Virus movement through PD is
probably an early infection event for many plant viruses [50,51]
and happens before virus-expressed fluorescent reporters can
be imaged. Microinjected fluorescently labelled TMV vRNA
was not detected in neighbouring cells even in MP transgenic
plants although the infection had spread, as shown by a virusexpressed FP [135]. Trafficked genomes were either recruited
Imaging plant virus infections
exclusively from the (unlabelled) pool of progeny genomes after
replication, or the amounts of fluorescence that were transported
across the cell–cell boundary were too small for detection. Other
studies have found cell–cell transport of viral RNA when MPs
were co-injected (e.g. [148]), possibly due to pre-formation of
movement complexes in vitro. Imaging of cell–cell transport
through PD is also hindered by difficulties in imaging the wallembedded PD, which is prone to the risk of optical artefacts due to
autofluorescence [64] and reflection [149]. Systemic transport in
the phloem is even more difficult to image due to the embedding
of the vasculature deep within the plant tissue. But perhaps
most challenging is imaging the release and uptake events that
occur from and into vector organisms, as direct visualization
would require bringing both plant and vector under the lens of
a microscope at the time of virus exchange. This remains an area
where the ingenuity of researchers is most strongly challenged.
FROM CELLS TO MOLECULES: BRIDGING THE IMAGING GAPS
Electron tomography
Similar to LM, EM has made significant advances in the
last two decades [14], including expansion into 3D imaging
through ET (reviewed in [13,14,16,150]). In ET, relatively thick
◦
specimens (100–500 nm) are tilted over typically +
−65 in 1–
2◦ increments along an axis perpendicular to the electron beam
during acquisition of 50–100 images. This set of images allows
computational 3D reconstruction with a z-resolution of 2–10 nm,
removing the limitation set by section thickness (∼ 20–100 nm)
of conventional EM. The digitized 3D model can then be reduced
to only selected structure outlines of interest. ET can thus enable
the analysis of 3D parameters such as volume, surface area or
connectivity of subcellular structures at unprecedented resolution.
Volumetric analysis of isolated macromolecular complexes by
cryo-ET even achieves sub-nanometer resolutions sufficient for
macromolecular structure determination, which is particularly
valuable when crystallization is problematic as in the case of
membrane proteins [151].
Electron tomographic studies have yielded remarkable insights
into the structure of animal viral replication sites (e.g. flock
house virus [152]; Figure 3a) and SARS (severe acute respiratory
syndrome)-coronavirus [153]. The flock house virus study
permitted calculations of replication site stoichiometry: virusinduced invaginations of the outer mitochondrial envelope
membrane each contain approx. 100 molecules of the replicative
transmembrane protein A, as well as an average of three RNA
replication templates, and remain connected to the cytoplasm
by a 10 nm membrane channel wide enough to permit export
of progeny RNA [152]. Such information is highly valuable
as it connects directly to recent biochemical insights into
how the oligomeric organization of viral RNA-dependent RNA
polymerase complexes assist in their overall function [154], which
may ultimately lead to the discovery of new antiviral drug targets.
ET has also been used to study animal virus entry [155,156],
uncoating [157], encapsidation [158] and exit events [159], and
cryo-ET has facilitated the study of complex, membranous virus
particles [160]. Most recently, successive sample abrasion with
a focused ion beam has been used as an alternative to a tilt
series for gaining 3D information in EM. This approach revealed
the connectivity of intracellular membraneous compartments
containing HIV particles with the plasma membrane [161].
For plant viruses, ET studies are still missing, although EMtomography has been very successfully applied to the study of
the plant endomembrane system and cytokinesis (e.g. [162,163]),
and was also used to study rice dwarf virus in its insect vector
31
[164,165]. It can be expected that plant viruses, also, will soon be
imaged within their plant hosts by ET.
For highly resolving EM approaches, high-pressure freezing
has become a standard technique that preserves sample
structure much better than traditional aldehyde fixation protocols,
especially in the case of membranous organelles [12]. Under
high pressure, samples up to a few hundred nanometers thick
can be frozen sufficiently fast to maintain cellular water
in an amorphous (vitrified) state, preventing tissue damage
by ice crystals. Frozen-hydrated samples can be sectioned
(cryo-ultramicrotomy; reviewed in [13,14]) and imaged directly
(cryo-EM), with resolutions up to 3–5 nm on cellular samples,
enough to identify known macromolecular complexes by their
shape [166,167]. However, unstained vitrified specimens provide
only phase contrast to generate detail and are also extremely
sensitive to the electron beam. This results in low signal/noise ratio
and makes it very difficult to pre-select a ROI, so that tomograms
sometimes have to be aquired ‘blindly’ [16]. Therefore alternative
or complementing approaches are required to increase contrast,
and permit molecular identification of cellular components.
The cellular water in high-pressure frozen samples can be
replaced with organic solvents and then embedding plastics
without thawing the sample (freeze substitution). Freeze
substitution permits the use of stains to increase contrast,
and embedded samples are stabilized for further sectioning
at room temperature as well as being less sensitive to the
electron beam although mostly retaining their preservation [168].
Truly specific labelling of proteins in ET can be achieved by
immunogold labelling and similar immunodetection of dsRNA
[153] or of bromo-UTP-labelled newly synthesized RNA [152],
is also possible. However, antibodies cannot be applied in
organic solvents or on frozen samples. Therefore high-pressure
frozen/freeze-substituted specimens commonly used for ET can
only be treated after embedding and sectioning, effectively
limiting immunogold labelling to the surface of the specimen.
Conversely, pre-embedding labelling gives greater sensitivity and
higher labelling levels [6], but requires chemical fixation. As a
compromise, several approaches have been developed to combine
ultrathin cryosectioning, which provides a better label/sample
volume ratio, with immunolabelling. This requires thawing and
therefore also chemical fixation at some stage. Fixation can
be done prior to freezing (Tokuyasu method), after thawing
or by thawing and rehydrating high-pressure frozen/freezesubstituted samples followed by re-freezing and cryosectioning
(see [169] for comparison). All of these approaches lose some
of the pristine sample preservation of direct cryo-EM/ET, and
chemical fixation also generally inactivates a proportion of the
accessible epitopes, preventing quantitative labelling [13,14]. As
a result, macromolecule-specific labelling techniques for EM/ET
currently fall short of complementing the available spatial imaging
precision. An EM-equivalent to FPs, i.e. a genetic tag than can
be directly detected throughout an EM sample would be highly
desirable [14].
CLEM (correlative light and electron microscopy)
and ‘super-resolution’
An alternative, or even preferable, approach would be to combine
the advantages of LM and EM (specific, quantitative protein
labelling and high resolution respectively) by correlative imaging
of the same sample (CLEM; reviewed in [15,16]). The promise of
CLEM has increased even further with the recent development
of various LM approaches that break the diffraction barrier,
long viewed as a limit to resolution in LM [9–11,170]. Because
c The Authors Journal compilation c 2010 Biochemical Society
32
Figure 3
J. Tilsner and K. J. Oparka
Bridging the imaging gaps
I. Extending EM to 3D by electron tomography. (a) FHV (flock house virus) induces invaginations of the outer mitochondrial membrane, where it replicates (i). Tomographic 3D reconstruction of
these invaginations (ii) shows them to be individual compartments, each of which is linked to the cytoplasm by a ∼ 10 nm channel (iii and iv). Using biochemical and structural information, the
potential packing arrangement of the replicative viral protein A within the spherule can be modelled (v) (reproduced from [152] under the Creative Commons Attribution License). II. Correlative light
‘super-resolution’ microscopy and EM. (b) Diffraction-limited TIRF (total internal reflection fluorescence) microscopy (i), ‘super-resolution’ PALM and transmission EM (iii) of the same cryosection
showing mitochondria. Overlay of PALM and EM images (iv). Reprinted from [173] with permission from AAAS. III. Combination of LM, EM and AFM (c) The ORF3 protein of groundnut rosette virus
localizes in cytoplasmic inclusions (i) that contain fibrillar material (ii). In EM cross-sections, the fibrillar material can be seen to consist of ring-like protein complexes encapsidating the viral RNA
(inset in ii). Fluorescence microscopy of ORF3–GFP (iii) and anti-fibrillarin immunofluorescence (iv) shows that ORF3 targets the nucleolus and leads to redistribution of nucleolar fibrillarin into the
cytoplasmic inclusions (v). In vitro , ORF3 and fibrillarin are both necessary to encapsidate viral RNA (vi–ix), but can form ring-like complexes in the absence of RNA (viii). These are similar to the
structures observed in infected tissue (ii, inset) and AFM reveals them to consist of alternating ORF3 and fibrillarin subunits (x) (reprinted by permission from Macmillian Publishers Ltd: EMBO
c 2007, reproduced from [38] c 2007 National Academy of Sciences, U.S.A., reproduced from [187] with permission from American Society for Microbiology, reprinted from [188]
Journal [37] c 2008 with permission from Elsevier). c-RNP, cytoplasmic RNP inclusion; DC, densely coated RNA; F, protein filaments; N, nucleus; No, nucleolus; V, vacuole; XB, cytoplasmic inclusion. Scale
bars: (a, panels i and ii), 100 nm; (a, panels iii and iv), 25 nm; (b), 1 μm; (c, panels i, iii, iv and v), 5 μm; (c, panel ii), 100 nm; (c, panel ii inset), 25 nm; (c, panels vi, vii, viii and ix), 100 nm; (c,
insets in panels vi, vii, viii and ix), 50 nm; (c, panel x), 20 nm.
light is subject to diffraction in all lens-based optical systems,
a fluorescent spot will appear as a PSF(point-spread function),
i.e. enlarged. Two fluorescent spots whose point spread functions
overlap cannot be resolved in a conventional light microscope
and according to the principle described by Ernst Abbe in 1873,
c The Authors Journal compilation c 2010 Biochemical Society
this limit to resolution is approximately half the wavelength
of the emitted light, ∼ 250 nm in the focal plane (x–y) and
∼ 500 nm along the optical axis (z) for fluorescence microscopy.
A number of different approaches now achieve resolutions
beyond the diffraction barrier and are collectively referred to
Imaging plant virus infections
as ‘super-resolution’ or ‘far-field’ microscopy or ‘nanoscopy’.
These include STED (stimulated emission depletion) microscopy,
which increases x–y resolution to tens of nanometers by focusing
a second laser beam on the excitation spot which is ring-shaped
and of an appropriate wavelength to quench excited fluorophores,
effectively restricting excitation to the dark centre of the
quenching laser [171]. SIM (structured illumination microscopy)
accesses subdiffraction information by moving a diffraction grid
over the sample and calculating additional spatial information
from the different diffraction patterns resulting from different grid
orientations [172]. Both STED and SIM are suitable for in vivo
imaging. PALM (photoactivation localization microscopy; [173]),
and STORM (stochastic optical reconstruction microscopy) is the
highest-resolving super-resolution technique [174], achieving up
to 5 nm lateral resolution. Imaging of individual fluorophores
without interference from overlapping point spread functions
allows precise calculation of the position of the actual fluorescent
spot within its PSF. In PALM/STORM, small numbers of
photoswitchable fluorophores are activated for each image,
localized and then bleached or switched off. This procedure
is repeated until the population of fluorescent molecules has
been exhausted and the collective image can be assembled
from the localizations of individual fluorophores. Reversibly
photoswitching pc-FPs such as Dronpa are particularly suited
for this approach [175]. The high resolution and the slow image
acquisition make PALM suitable mostly for fixed specimens. In
all these techniques, additional modifications are necessary to
achieve super-resolution also along the light path in the axial
dimension. Various approaches have been developed including
the use of two opposing objectives (4Pi microscopy), z-dependent
lateral distortion of the image or axial interference patterns, and
z-resolutions of approx. 50 nm can be achieved (reviewed in
[10,170]). In vivo 4Pi microscopy has been correlated with EM
to track transport of cargo molecules through single Golgi stacks
and PALM has been combined with cryo-EM [173] (Figure 3b).
The power of such correlative approaches is evident from the
achievable labelling density: Betzig et al. [173] counted >5500
fluorophores in a subset of mitochondria in a single PALM
cryosection, whereas only 10–20 labels/section could be expected
in a comparative immunogold experiment [86].
To image the same sample in CLEM, either live-cell imaging
has to be carried out before fixation or the same fixed specimen has to be used in both LM and EM. The first approach
makes full use of the benefits of LM, and ideally would
allow the cessation of cellular processes at a precise moment
of interest. Cryosectioning is ideal for the preservation of
both fluorophores and sample ultrastructure, but continuous
maintenance of samples at vitreous temperatures (<−140 ◦C)
is technically extremely challenging [16]. Less demanding is
spatial matching of chemically fixed samples in both microscopies
for which the first commercial setups are on the horizon
(reviewed in [15]). Unfortunately, in fixed samples, the use
of the fluorescent markers becomes problematic. Chemical
fixatives denature proteins, and even though GFP-derivatives are
very resilient and often retain fluorescence after mild fixation,
signal intensity is usually significantly diminished, making this
procedure unsuitable for low-abundance FP-fusions. If the sample
is embedded in polymers for sectioning, additional problems
may arise because conventional FPs require molecular oxygen
for fluorescence. Such problems can potentially be solved by
use of iLOV, which does not require oxygen (but does require
presence of FAD) [63,176], or fluorescent labelling during/after
embedding, e.g. with tag–probe labelling systems or using
fluorescent antibodies. The best currently available tool for this
purpose, which also comes closest to a ‘GFP for EM’, is the
33
photo-oxidative precipitation of DAB (diaminobenzidine) by
fluorophore bleaching [15,177,178]. Oxygen radicals generated in
the vicinity of fluorophores during photobleaching oxidize DAB,
which forms a granular polymer that can be osmium-stained,
rendering it EM-visible. This technique has been demonstrated for
FPs [178,179] as well as tetracysteine-/ReAsH-labelled proteins
[180] and various fluorescent dyes [15,177,178].
Also suitable for direct LM/EM correlation are Qdot (quantumdot)-labelled antibodies. Qdots are fluorescent nanocrystals which
neither bleach [64] nor require molecular oxygen. Their spectral
properties are defined by their size and they are large enough and
sufficiently electron-dense to be directly visible in EM [181]. Use
of Qdots for live-cell imaging has been hampered by their large
size and incompatibility with biological molecules. However,
CLEM may be the area where their full utility can be realized
[15]. It can be expected that the combined imaging power of
super-resolution LM and EM will provide many new insights into
previously inaccessible aspects of plant virus infections in the
near future.
Combined EM and AFM
At the far end of the resolution scale, EM can also be combined
with AFM, which can achieve true atomic resolution [182].
In AFM, a very fine needle tip is used to scan the surface
of the sample, either by touching it directly (contact mode)
or by remaining at a small distance utilizing electromagnetic
interactions (non-contact mode). Needle movements are measured
with high accuracy and allow 3D reconstruction of the sample
surface. The most significant contribution of AFM to plant
virology so far has been in the study of isolated virions and viral
RNPs (ribonucleoprotein particles).
Unlike EM, AFM does not require staining, which can obscure
small features and is therefore useful for discovering assymetric
structures of viral RNPs which can then be identified by immunoEM [183]. Interestingly, assymetric binding of viral proteins to
only one end of virions or viral ribonucleoprotein complexes has
been found for a number of very different viruses (e.g. [84,183–
186]) and may represent a general principle by which virions
functionally interact with host and/or vector organisms. AFM
and immuno-EM were used to investigate the destablizing effect
on the PVX virion of binding by the TGB1 MP to its 5 end
[84,184]. These experiments established that binding of TGB1
is one way to render the encapsidated viral RNA translationcompetent and also identified an in vitro-assembled complex of
viral RNA partially encapsidated in CP, termed a ‘single-tailed
particle’ as the potential transport form of this virus [84]. Lim
et al. [48] recently isolated viral RNPs from plants infected with a
mutant form of barley stripe mosaic virus lacking the coat protein.
The absence of CP prevents formation of true virions and these
RNPs, which contained the TGB1 MP and positive sense genomic
and subgenomic ssRNAs, were assumed to be viral movement
complexes. Low amounts of unindentified high-molecular-mass
protein components were also found, but the minor MPs TGB2
and 3 were not detectable, despite heterologous interactions
between the TGBs [48]. Combined AFM and immuno-EM
imaging of the complexes might be a useful approach to elucidate
the potential presence, identity and position in the RNP of minor
protein components. Similar approaches could be adopted for
other viruses that do not require their coat protein for local
movement, e.g. TMV.
In the case of the recruitment of nucleolar fibrillarin by the
groundnut rosette virus ORF3 protein [37,38], described above,
combined EM and AFM showed that fibrillarin forms part
of the ring-like complexes that protect the viral RNA during
c The Authors Journal compilation c 2010 Biochemical Society
34
J. Tilsner and K. J. Oparka
long-distance transport [38,187,188]. This was the first direct
demonstration of a plant host protein participating in a viral
movement complex (Figure 3c). The study of viral RNPs by
combined AFM and EM has provided some of the deepest insights
into the elusive nature of putative plant viral movement complexes
and will become even more valuable when more structural data on
MPs become available. Collectively, the combined power of the
individual and correlative imaging approaches now at the disposal
of plant virologists hold the promise that we will be able to gain
a truly molecular understanding of all stages of the viral infection
cycle within the not-so-distant future.
ACKNOWLEDGEMENTS
We apologize to all collegues whose work we were unable to cite due to space limitations.
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Received 12 March 2010/20 May 2010; accepted 1 June 2010
Published on the Internet 28 July 2010, doi:10.1042/BJ20100372
c The Authors Journal compilation c 2010 Biochemical Society