To study the
properties of individual PP1 isoforms in living cells, including
their localization and interaction with targeting subunits, we have
taken advantage of the in vivo approach of fusing PP1 to fluorescent
reporter molecules, in this case chromatic variants of the green
fluorescent protein (GFP) derived from the jellyfish Aequorea victoria.
This method effectively “tags” the intracellular pool
of the protein and allows analyses of dynamic properties in living
cells. Their changing spatio-temporal distributions can be monitored
both throughout the cell cycle and following cellular perturbations
by time-lapse fluorescence microscopy, and turnover rates of intracellular
pools of the protein calculated by fluorescence recovery after photobleaching
(FRAP). Interactions with targeting subunits can be visualized in
vivo by fluorescence resonance energy transfer (FRET), using techniques
as sensitized emission, acceptor photobleaching, or fluorescence
lifetime imaging (FLIM/FRET).
|Live Cell Time-Lapse Imaging
For imaging live cells in glass bottom dishes or POC chambers,
replace the standard growth medium with Phenol Red-free medium supplemented
with FBS and penicillin-streptomycin. If using an environmental
chamber supplied with 5% CO2, pH will be maintained. If not, add
20 mM HEPES to the medium as a buffer. Alternatively, a custom-made
CO2-independent Phenol Red-free medium can be used, supplemented
with FBS and penicillin-streptomycin. This medium does not require
additional buffering. DNA can be stained in live cells, with the
cells remaining viable and progressing through the cell cycle. To
do this, pre-treat cells for 30 min with Hoechst No. 33342 dye.
It is important to note that this dye cannot be used with ECFP-tagged
proteins since the two cannot be spectrally resolved.
When imaging cells through the cell cycle, prophase cells can be
found by their characteristic condensed DNA pattern. Minimize the
amount of light the cells are subjected to during imaging by keeping
exposure times as low as possible while maintaining adequate resolution.
Hoechst No. 33342 can be imaged using a DAPI filter set, while EGFP
can be imaged using a FITC filter set. EYFP and ECFP require special
filter sets. Recently, we have used EGFP in combination with mCherry,
which can be separated using a FITC/TRITC filter set.
|Time-lapse imaging of FP-PP1. HeLa cells stably expressing
FP-PP1γ and stained with the cell permeable DNA dye Hoechst
No. 33342 were imaged as they progressed through mitosis. Images
were taken every 3 minutes using low light levels and short
exposure times to minimize photodamage. This cell starts in
prophase, with the DNA condensing and FP-PP1γ still visible
within nucleoli. Within 12 minutes it has progressed to metaphase,
at which point FP-PP1γ is primarily diffuse but also shows
an accumulation at kinetochores. As the cell progresses through
anaphase (at 15-18 minutes), FP-PP1γ shows a relocalization
to chromatin, where it remains until it starts to re-accumulate
in nucleoli at late telophase (just visible at 27 minutes).
|FRAP (fluorescence recovery
Although time-lapse imaging can only show the steady-state distribution
of a protein over time, FRAP analyses can reveal its kinetic properties,
such as whether it’s free to diffuse or immobilized to a scaffold,
and if it’s exchanging between particular compartments and
at what rate. In short, bleaching a fluorescent pool of protein
in one region of the cell using a high-intensity laser pulse renders
that protein “invisible”, and recovery can then be monitored
as the unbleached molecules from neighboring regions of the cell
move into the bleached region.
FRAP experiments are generally performed on cells expressing GFP-
or EGFP-tagged proteins (EYFP shows a small but significant amount
of spontaneous recovery of the molecule from photobleaching, which
complicates the interpretation of FRAP results), and can be done
using either laser scanning microscopes or widefield fluorescence
microscopes equipped with an external laser for photobleaching.
When analyzing proteins with very fast turnover rates, the microscope
system used must offer the appropriate temporal resolution (i.e.
image rapidly enough to capture early events).
To perform a FRAP experiment, take 2-3 images of the cell prior
to photobleaching and then bleach a region of interest to approximately
50% of its original intensity, acquiring images over time after
the photobleach period to monitor recovery of fluorescence signal
within the bleached region. If recovery is observed, it indicates
that the fluorescently-tagged proteins are mobile, and the rate
of recovery is therefore an indication of the speed at which they’re
moving. For comparison, the same photobleaching experiment can be
conducted on fixed cells, in which no recovery is observed.
For qualitative FRAP, it may be enough to simply plot fluorescence
intensity over time (known as the recovery curve), as a fraction
of the initial fluorescence prior to photobleaching. Different pools
of the same protein can be compared for relative mobility, or the
same pool of protein can be assessed before and after a particular
For quantitative FRAP, the most commonly published results are the
mobile fraction (fraction of fluorescent molecules that are free
to move within the bleached region) and the half-time for recovery
of fluorescence (t1/2), which is the time required for half of the
mobile fraction to recover. The t1/2 can then be used to calculate
the diffusion coefficient (for reference see: Carrero G., McDonald
D., Crawford E., de Vries G. and Hendzel, M.J. (2003) Using FRAP
and mathematical modeling to determine the in vivo kinetics of nuclear
proteins. Methods. 1:14-28.)
|Example of a FRAP experiment in which the
turnover of three different cellular pools of EGFP-PP1γ
were compared (cytoplasmic vs. nucleoplasmic vs. nucleolar).
A small circular region (hashed circle indicated by arrow) was
photobleached in a region of the cell and recovery of fluorescent
signal within that region monitored over time. By plotting recovery
of fluorescence intensity versus time after photobleaching ,
half-times of recovery can be calculated for each pool of FP-PP1γ.
Cytoplasmic FP-PP1γ shows the fastest recovery, with a
t1/2 of approximately 1 sec (comparable to that for free GFP).
Nucleoplasmic FP-PP1γ shows a slower recovery rate (t1/2
= 2 sec), as does nucleolar FP-PP1γ (t1/2 = 9 sec), but
these rates still indicate a rapid flux of PP1 through these
|FRET (fluorescence resonance
Interactions between proteins, such as PP1 and a targeting subunit,
can be shown in vivo using various FRET measurement techniques.
The basic principle of FRET is the transfer of energy from an excited
donor fluorophore to an acceptor fluorophore in close proximity.
FRET is strongly dependent on the distance between donor and acceptor,
falling off with the sixth power of the distance between the two.
Because of this, FRET can only occur when the proteins are within
1-10 nm of each other and in the proper orientation. A simplified
diagram is presented here, which demonstrates how excitation of
the donor sensitizes emission from the acceptor that ordinarily
would not occur. FRET can therefore be detected as sensitized emission
of the acceptor.
Energy transfer from donor to acceptor depletes or “quenches”
the excited state population of the donor, and FRET will therefore
reduce the fluorescence intensity of the donor. Photobleaching the
acceptor to relieve this quenching of the donor (termed “acceptor
photobleaching”) offers another option for detecting FRET
in vivo. FRET-induced donor quenching is also observed as a decrease
in the donor’s fluorescence lifetime, which is the average
time that a molecule spends in the excited state before emitting
a photon and returning to the ground state. Comparison of donor
lifetime in the presence and absence of acceptor is the last FRET
method presented here, termed the FLIM/FRET technique. Key advantages
of this approach are the independence of measurement on probe concentration,
and the use of infrared excitation wavelengths which are less damaging
to cells. For all three of these FRET techniques, the efficiency
of energy transfer may be used as a molecular ruler to determine
the scale of a particular interaction (for reference see: Stryer,
L. (1978) Fluorescence energy transfer as a spectroscopic ruler.
Annu. Rev. Biochem. 47, 819-846).
|Measuring FRET between PP1
and a targeting subunit by sensitized emission
This approach requires independent control of emission and excitation
filters. Measurements are taken using a combination of three filter
sets, ECFP (excite 436 nm, emit 470 nm), EYFP (excite 514 nm, emit
528 nm) and what we refer to as the FRET channel (excite 436 nm,
emit 528 nm). The principle of FRET is detailed in this cartoon.
For the experimental condition cells should be co-transfected with
EYFP-PP1 and ECFP-targeting subunit (or the inverse, ECFP-PP1 and
EYFP-targeting subunit. For control measurements, transfect some
cells with the EYFP construct alone, and some with the ECFP construct
alone. As a negative control, co-transfect cells with PP1 and a
mutant version of the targeting subunit that cannot bind PP1 (e.g.
mutation of one or both hydrophobic residues in the RVXF motif).
An alternate control is the ECFP-tagged protein co-expressed with
an EYFP-tagged protein that shows the same localization but does
not interact with it.
To correct for spectral bleed-through, first examine cells expressing
either donor alone or acceptor alone with each of the three filter
sets. For example, using our system, ECFP-NIPP1 alone gives a strong
signal in the ECFP channel and no signal in the EYFP channel, but
it does show significant spectral bleed-through into the FRET channel
(~70% of the signal measured in the ECFP channel). EYFP-PP1 alone
gives a strong signal in the EYFP channel and no signal in the ECFP
channel, but also shows spectral bleed-through into the FRET channel
(~20% of the signal measured in the EYFP channel).
Measure FRET in cells expressing both PP1 and targeting subunit
by exciting at the ECFP wavelength (436 nm) and detecting at the
EYFP emission wavelength (528 nm). The following equation is used
for the correction of spectral bleed-through, in which FRET(N) is
Net Energy Transfer:
FRET(N)= FRET signal - α(donor signal)
- β(acceptor signal)
*For this equation, α and β are determined by imaging
the cells expressing each fusion protein on its own (0.7 for
ECFP-NIPP1 and 0.2 for EYFP-PP1).
Certain software programs, such as the SoftWorx analysis software
used with the DeltaVision system, allow image subtraction, to obtain
a final FRET image showing the signal remaining in the FRET channel
following this correction.
The same analysis should then be applied to data collected from
cells expressing PP1 and a mutant targeting subunit that cannot
interact with it, or alternatively two proteins that show the same
localization but do not interact. After the data have been corrected
for spectral bleed-through, there should be no signal remaining
in the FRET channel.
|Positive control is shown on the left (FRET
between PP1 and wild type NIPP1), and negative control is shown
on the right (no FRET between PP1 and NIPP1 in which the PP1
binding site has been mutated). Note that in the latter case
the mutant NIPP1 cannot retarget PP1 out of the nucleolus because
it does not interact with it.
|Measuring FRET between PP1
and a targeting subunit by acceptor photobleaching
This approach, which is based on the increase in donor (ECFP)
signal when FRET is disrupted by photobleaching the acceptor (EYFP)
molecule, requires a laser line at approximately 532 nm to photobleach
EYFP (without bleaching ECFP) and appropriate excitation and emission
filter sets to monitor ECFP fluorescence before and after this bleaching.
It can be done using either a laser scanning microscope such asa
Zeiss LSM 51 510 or a widefield fluorescence microscope equipped
with an external laser, such as the DeltaVision Spectris. The DeltaVision
system offers slightly greater flexibility due to its improved temporal
resolution, since the increased donor signal is only apparent until
the photobleached pool of acceptor recovers and once again quenches
the signal. If the protein happens to turn over rapidly, as does
PP1, then the unquenched donor signal will only be observed at very
early time points. This can be compensated for when using a system
with slower post-bleach imaging resolution by photobleaching a larger
area of the cell, to ensure that acceptor recovery does not mask
the unquenched donor signal in the region of interest.
Cells should be co-transfected with EYFP-PP1 and ECFP-targeting
subunit, or alternatively ECFP-PP1 and EYFP-targeting subunit. After
obtaining several pre-bleach images of both the donor and the acceptor
proteins, photobleach the acceptor in a region of the cell using
the appropriate settings for the system used (e.g. 100% laser power
and several iterations). After bleaching, collect images at the
desired time intervals, using the same settings used to obtain the
Analyze the data utilizing the image analysis tools included in
the imaging system’s software. Most systems now include FRAP
analysis tools in their software packages. Spreadsheet and biostatistics
programs such as Microsoft Excel (Microsoft Corp., California, USA)
and GraphPad Prism (GraphPad Software Inc., California, USA) are
also useful tools for the analysis of photobleaching data. In addition
to the bleached region, include a region in the non-bleached portion
of the same cell or in a neighboring cell in the data analysis as
a control for bleaching due to imaging. A region of background fluorescence
should also be defined outside the cell, and subtracted from both
the bleached and control regions.
The FRET efficiency can be calculated
using the following formula:
FRET Efficiency E = (ID(post) –
* ID(pre) and ID(post) are donor intensity
before and after photobleaching, respectively
Bleach Efficiency B = (IA(pre) –
Corrected FRET Efficiency: (E/B) x 100%
Using these equations, the ECFP-PP1/EYFP-NIPP1 interaction
shown here has a FRET efficiency of 10.2%.
between PP1 and a targeting subunit by FLIM/FRET
To briefly summarize the principle of lifetime imaging, the fluorescence
of organic molecules is not only characterized by their excitation
and emission spectra, but also by their lifetimes. When a fluorophore
absorbs a photon it goes into the excited state and returns to the
ground state by emitting a fluorescence photon, converting the energy
internally, or by transferring the energy to the environment. Although
fluorescence lifetime imaging systems can seem overly complex at
first, in reality the basics are fairly easy to grasp (see Fig.
4). We use the BioRad Radiance Multiphoton Imaging System, which
allows us to obtain both intensity and lifetime images for the fluorescent
proteins of interest in live cells.
Cells should be co-transfected with either ECFP-targeting subunit
and EYFP-PP1 or ECFP-PP1 and EYFP-targeting subunit. For control
measurements, transfect cells with the ECFP construct alone (to
measure the lifetime of unquenched ECFP), or with the ECFP construct
and an EYFP-tagged protein that shows the same localization but
does not interact with it. Excite ECFP at a wavelength of 840 nm,
allowing the external detectors to collect the fluorescent emission
and calculate lifetimes.
In the figure below, lifetime maps have been plotted (i.e. each
pixel is color coded with a lifetime value to build up an image
of differences in lifetime throughout the sample) for ECFP-NIPP1
in the presence of either EYFP-PP1 or EYFP-U1A. They are shown next
to corresponding intensity images. Quenching of the donor (ECFP-NIPP1)
causes a shift to a shorter lifetime, and FRET is thus observed
as the appearance of a second, “quenched” lifetime (~1.6
ns, compared to the unquenched lifetime of ~1.9 ns). In the presence
of EYFP-U1A, which shows the same localization pattern but does
not interact with ECFP-NIPP1, only a single, unquenched lifetime
of ~1.9 ns is observed. The quenched and unquenched lifetimes can
also be color-coded to demonstrate more clearly where FRET is occurring
within the cell. In this example, although NIPP1 is found throughout
the cell nucleus, the predominant FRET signal is observed at nuclear
speckles, where mRNA splicing factors are known to accumulate, suggesting
a role for the complex in the regulation of pre-mRNA splicing.
The FRET efficiency can be calculated
using the following formula:
FRET Efficiency E = 1-(τDA/τD)
* τDA and τD are donor lifetime
in the presence and absence of acceptor, respectively
Using this FRET efficiency, the
distance between the donor and acceptor can also be calculated:
Rdonor/acceptor = R0((1/E)-1)1/6
* R0 = Forster distance, which is
50Å for the CFP/YFP pair
Using these equations, the ECFP-PP1/EYFP-NIPP1 interaction
shown here has a FRET efficiency of 15.8%, which translates
into a donor/acceptor distance of approximately 66Å.
I have written a book chapter that details the use of fluorescent
PP1 isoforms in microscopy experiments. This chapter includes detailed
protocols and information for sourcing reagents:
L., Chusainow, J., Lam, Y.W., Swift, S. and Lamond,
A.I. Visualization of intracellular PP1 targeting through
transiently and stably expressed fluorescent protein fusions.
Methods in Molecular Biology. Vol.. 365:133-54; Protein Phosphatase
Protocols, Ed. G. Moorhead, Humana Press, 2006.
I have also written a series of basic microscopy review articles
with Dr. Sam Swift, manager of the light and electron microscopy facility
at the University of Dundee:
Swift, S. and Trinkle-Mulcahy, L. To bin or not to bin: Balancing the trade-off between signal intensity, spatial and time resolution in biological imaging In Focus 26:5-14.
L. and Swift, S. Caught on camera with another protein -
Just good friends or something more? A Guide to Localization, Colocalization
and Interaction on the Light Microscope. In Focus (Proceedings of
the Royal Microscopy Society), 8:4-15, 2007.
S.R., Thomson, C., Appleton, P., Lam, Y.W. and L. Trinkle-Mulcahy.
Basic principles deconvolution microscopy. Proc. Royal Mic. Soc.,
S.R. and L. Trinkle-Mulcahy. Basic principles
of FRAP, FLIM and FRET. Proc. Royal Mic. Soc., 39:3-10, 2004.