Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)
Live CLEM imaging: an application for yeast cells
Haruhiko Asakawa1, Yasushi Hiraoka1,2, 3, and Tokuko Haraguchi1,2, 3
1
Graduate School of Frontier Biosciences, Osaka University, Suita 565-0871, Japan
Advanced ICT Research Institute Kobe, National Institute of Information and Communications Technology, 588-2
Iwaoka, Iwaoka-cho, Nishi-ku, Kobe 651-2492, Japan
3
Graduate School of Science, Osaka University, Toyonaka 560-0043, Japan
2
Correlative light and electron microscopy (CLEM) is a useful tool for investigating cellular structures at high-resolution.
We have extended this technology and developed a new method in which CLEM is combined with live-cell fluorescence
imaging, “Live CLEM”. With this method, living cells are first examined by fluorescence microscopy then fixed in situ,
and finally the same cells are subjected to electron microscopy. This simple yet powerful method combines temporal and
high resolution spatial information. However, due to technical difficulties in immobilizing non-adherent target cells during
sample preparation, this method has usually been applied to the study of adherent cells. Here we report a method of live
CLEM that can be applied to yeast cells. Since yeast are a powerful tool in molecular genetics, application of this method
to the study of yeast should greatly advance our understanding of the molecular basis of the function of cellular structures.
Keywords Fluorescence microscopy; electron microscopy; live-cell imaging; correlative light-electron microscopy;
CLEM
1. Introduction
Fluorescence microscopy (FM) is a powerful tool for observing specific molecular components in living cells, but it can
only provide relatively low-resolution imaging. In contrast, electron microscopy (EM) provides high-resolution imaging
of cellular structures, but it cannot provide temporal information in living cells. To overcome these inherent problems,
we have developed a method of combining EM imaging with live-cell FM imaging, “Live CLEM” (Live cell imaging
associating Correlative Light and Electron Microscopy). Live CLEM is a 2-stage method in which the dynamic
behavior of specific molecules of interest in a living cell are first observed using FM and then cellular structures, such
as organelles and membranes, in the same cell are observed using EM. Following image acquisition, FM and EM
images are compared to enable the fluorescent images to be correlated with the high-resolution images of cellular
structures obtained using EM. This method enables analysis of dynamic events involving specific molecules of interest
in the context of specific cellular structures. Thus, it is a powerful tool for examining dynamic and complex biological
events such as nuclear envelope reformation and autophagosome formation in mammalian cells [1, 2]. To date, due to
technical difficulties associated with the movement/drift of non-adherent cells in the medium, particularly during
preparation of the sample for EM following the acquisition of FM images, the method has usually been applied to the
study of adherent cells. We have developed a method involving concanavalin A, a lectin (sugar-binding protein), to
immobilize yeast cells for Live CLEM imaging. Here we describe in detail this method of Live CLEM imaging:
combining Live CLEM imaging with yeast genetics creates a powerful genetic tool which can be used to investigate the
function of cellular structures at the molecular level.
2. Materials and methods
2.1
Fluorescence microscope set-up for live cell imaging of yeast cells
Several fluorescence microscope systems capable of live-cell imaging are commercially available. With particular
relevance for live CLEM imaging, a system capable of obtaining z-stack images is highly recommended. In addition,
deconvolution of the 3D images improves FM images by removing out-of-focus images [3]. In our laboratory, the
DeltaVision system (Applied Precision, Inc., Seattle, USA) based on an Olympus wide-field fluorescence microscope
IX70 (Olympus Corp., Tokyo, Japan) is used. Fluorescence images are obtained using the interline CoolSNAP HQ2
CCD camera (Photometrics, Tucson, USA) as an image detector through an oil-immersion objective lens (Plan Apo 60
×, NA = 1.4, Olympus). The softWoRx® software equipped with the microscope system is used for deconvolution.
During live observation, the microscope is kept at a desired temperature (26-30 °C ±1°C) in a temperature-controlled
room, and the computer and other control units are placed outside the room and control the microscope remotely, as
described previously [4]. Alternatively, various types of temperature control equipment are available commercially. The
choice of such equipment requires careful consideration if accurate temperature control during observation is to be
achieved [5].
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2.2
Fluorescence live cell imaging for CLEM
In general, living yeast cells are observed by either of two methods: sandwiching the cells between glass coverslips, or
by mounting the cells on a glass-bottom culture dish (a plastic culture dish with a glass coverslip attached to a round
opening in the bottom of the dish) [6]. Of the two methods, the glass-bottom dish approach is often used owing to the
convenience it offers for long-duration observations or for experiments requiring exchange of the bathing medium (Fig.
1): in this case, an inverse microscope, rather than an upright microscope as is typically employed in yeast laboratories,
should be used. For Live CLEM imaging, in addition to the equipment required for live cell imaging, a means of
identifying specific cells is needed. We use a gridded glass-bottom dish (a plastic culture dish with a gridded coverslip,
instead of a regular coverslip, attached to the opening in the bottom of the dish). Several types of such gridded
glass-bottom dishes, that are suitable for Live CLEM for yeast cells, are commercially available, and the features of
some of these are summarized in Table1.
2.2.1
Preparation of yeast cells
- Isolate single colonies by streaking a strain harboring the GFP-fusion gene of interest on an agar plate containing
complete medium. Use a selective culture medium when auxotrophic selection is applied. The composition of each
medium can be found in the literature [7, 8].
- Incubate at 30 °C until colonies are formed (typically for 3-4 days).
- Pick a single colony using a toothpick and suspend in 5-10 mL complete liquid culture medium.
- Incubate at 30 °C with vigorous shaking until the culture reaches logarithmic phase (0.2-1× 107 cells/mL).
- Collect cells by centrifugation at 3000 rpm (1200 × g) for 5 min.
- Suspend cells in an appropriate synthetic medium. Use an optically clear and colorless medium as a turbid medium
disturbs microscopic observation.
2.2.2
Immobilization of cell specimens for microscopic observation
For Live CLEM imaging, a cell is traced during live cell imaging and that cell is analyzed by both FM and EM.
However, yeast cells are hard to trace during live cell imaging because they are non-adherent and tend to move easily
(e.g., due to stage movement, convection flow of culture medium, contact and pushing of neighboring cells during cell
growth, etc). To immobilize the yeast cells, we coat the glass surface of the glass-bottom dish with concanavalin A or
lectin before use. For the fission yeast Schizosaccharomyces pombe, either concanavalin A or soybean lectin is used,
while for the budding yeast Saccharomyces cerevisiae, concanavalin A is used.
- Drop and spread 50 μL of 2 mg/mL aqueous concanavalin A or 0.2 mg/mL aqueous soybean lectin onto the glass
surface of the glass-bottom dish.
Fig. 1 Preparation for live cell imaging. a, An example of
a plastic culture dish with a gridded coverslip, instead of a
regular coverslip, attached to the opening in the bottom of
the dish (ibidi plastic bottom dish 35 mm grid-500;
product number 81161). b, An example of the grid pattern
for an ibidi 35 mm dish with a 500 μm grid. c-e,
Preparation of culture dish for live cell imaging. c, A piece
of moistened paper is placed in the dish for long-duration
observations, and a droplet containing the cell suspension
is placed on the grid. d, The lid is placed on the dish and
sealed with Parafilm. e, Fluorescence microscope stage
setup. An inverted microscope is used for the study of
cells in a glass-bottom dish.
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Current Microscopy Contributions to Advances in Science and Technology (A. Méndez-Vilas, Ed.)
Table 1 Features of commercially available gridded culture dishes
Manufact
urer, URL
Ibidi,
http://ww
w.ibidi.co
m/
Product
no.
81161
81148
81168
Features
Comments
- Special plastic-bottom dish,
- Plastic parts, except for the lid, are
resistant to acetone
- Thickness of the plastic bottom is
equivalent to a No. 1.5 glass
coverslip*
- Optically transparent
- Very low or no fluorescence
background
- Gas permeable
- Grid size 500 μm
- Glass-bottom dish
- Plastic parts, except for the lid, are
resistant to acetone
- Coverslip thickness, No. 1.5*
- Grid size 50 μm
- Glass-bottom dish
- Plastic parts, except for a lid, are
resistant to acetone
- Coverslips thickness, No. 1.5*
- Grid size 500 μm
- Glass-bottom dish
- Sensitive to acetone
- Coverslip thickness, No. 2*
- Grid size 600 μm,
This plastic-bottom dish has very low background
fluorescence. Thus, fluorescence live-cell imaging of
yeast cells can be done successfully. However, taking
accurate fluorescence 3D images is relatively difficult
because the plastic bottom is relatively elastic and tends
to move toward a lens when changing the focus of a
fluorescence microscope. This elastic nature of the
plastic bottom may also cause a problem with EM
observation because it provides the epoxy block with
wavy surface. Removal of the epoxy block from the
dish is easy.
Removal of the epoxy block from the dish is relatively
difficult. If the dish is broken and glass pieces are left
on the surface of the epoxy block, use hydrofluoric acid
to remove the residual glass, as described in the text.
Removal of the epoxy block from the dish is relatively
difficult. If the dish is broken and glass pieces are left
on the surface of the epoxy block, use hydrofluoric acid
to remove the residual glass, as described in the text.
MatTech, P35G-2This dish can be used if the duration of the acetone
http://glas 14treatment is limited to 10 min or less.
s-bottom- CGRD
dishes.co
m/
*The thickness number is defined as follows: No. 1.5, 0.16-0.19 mm; No.2, 0.19-0.23 mm.
- After one minute, remove excess concanavalin A or soybean lectin solution and dry for several hours, drying
overnight is best.
- Place 50 μL of the cell suspension (cell density should be approximately 5000 cells per 1 mm2 glass surface) onto the
coated glass-bottom dish (Fig. 1c and e).
- To avoid desiccation during observations longer than several hours, place a piece of paper moistened with distilled
water in the culture dish (Fig. 1c) and/or place a lid on the dish and seal it with Parafilm (Fig. 1d).
2.2.3
Time-lapse observation
- Place the dish containing live cell specimens on the microscope stage and leave it for 30 min before image data
acquisition to allow the cells to settle and for the observation temperature to be reached.
- Run image acquisition software to obtain time-lapse image data.
2.3
High-resolution fluorescence 3D imaging of a fixed specimen
Acquisition of high-resolution fluorescence 3D images of a fixed target cell is helpful for finding the target cell or
subcellular structures of interest during EM observation. For this purpose, after fixing the cells at an appropriate time of
interest during the time-lapse observation, we obtain their fluorescence images immediately after addition of fixative:
- Remove the culture medium using a pipette.
- Add a fixative solution (approximately 50 μl of 2% glutaraldehyde (Polysciences Inc., Warrington, USA) in 0.1 M
sodium phosphate buffer (pH 7.2)).
- After one minute, carefully remove the fixative.
- Add fresh fixative solution.
- Obtain three-dimensional fluorescence images of the cell of interest (ideally, 20-30 focal planes at 0.2 μm intervals).
A washing step to remove fixative is not required for image acquisition when image acquisition is performed within 510 min after addition of the fixative.
- Obtain bright-field images at low magnification. Alternatively, make drawings indicating cell shapes, directions, and
relative positions of cells and grids. These images are important for later identification of the cell of interest fixed in an
epoxy block (see 2.4.3).
- Incubate for 2 hrs at 4°C.
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2.4
2.4.1
Electron microscopy
Fixation and embedding of cells for electron microscopy
- Post-fix cells for 2 hours with an aqueous solution of 1.2% KMnO4.
- Wash briefly with distilled water eight times, and sequentially dehydrate by replacing the water with increasing
concentrations of ethanol (50, 70, 80, 90, 95, 99.5, and 100%) at 10 min intervals.
- For embedding, incubate cells sequentially in acetone for 10 min, 50% (v/v) Epon812 (TAAB, Berkshire, UK) in
acetone for 1 hour, 66.7% Epon812 in acetone for 1 hour, and 100% Epon812 overnight. For this step, because dish lids
are usually not resistant to organic solvents such as acetone, remove the dish lid and use home-use polyvinylidene wrap
instead.
- Incubate in fresh 100% Epon812 for 3-4 hours.
- Physically fix the gridded glass-bottom dish onto a glass slide by dropping a few drops of 100% Epon812 onto the
glass slide (Fig. 2a) and placing the dish on it (Fig. 2b). This step makes it easier to strip off a sample block from the
dish, as shown in the next section (2.4.2).
- To make a block of epoxy resin containing the cell specimens, fill an embedding capsule with 100% Epon812 (Fig.
2c), and position it upside down on the area that contains the cell of interest (Fig. 2d).
- Incubate the resin at 60 °C for 24-48 hrs for polymerization.
2.4.2
Removal of sample blocks from a dish
- Heat the bottom of the dish using a spirit lamp (Fig. 2e).
- Strip the embedding capsule from the glass-bottomed dish using pincers (Fig. 2f). The grid is replicated on the surface
of epoxy resin block (Fig. 2g and h).
- If stripping is difficult, try soaking the glass slide and dish in a liquid nitrogen bath. The embedding capsule will come
away from the glass bottom because of the differences in the shrinking properties of the glass bottom and the epoxy
resin.
- Using a razor blade, cut and remove the embedding capsule.
- If the glass bottom is broken in the stripping process, hydrofluoric acid can be used to dissolve the residual glass
pieces of the glass-bottom dish: Pour the hydrofluoric acid into a small plastic tube (don’t use a glass tube! we usually
use a 48-well plastic plate), and, after any plastic parts are removed, soak the block with the glass fragments facing
upwards. After 20 min, wash the block several times with water, and observe using a microscope to confirm that the
Fig. 2 Sample embedding. a, A glass slide with a droplet of
epoxy solution. b, Mounting the dish on the slide glass. c, An
embedding capsule filled with resin. The lid has been
removed. d, The embedding capsule filled with resin is placed
on the grid upside down. e, The bottom of the glass-bottom
dish is gently warmed with a spirit lamp. f, The embedding
capsule is removed from the dish using pincers. g, The
embedded sample is removed from the coverslip with the grid
pattern clearly replicated. h, Schematic of embedding and
replica of grid pattern. The grid facing the cell suspension in
the culture dish is replicated in the epoxy resin upon
embedding. The replicated grid pattern serves as a landmark
to find target cells.
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glass has dissolved. If this is the case, air-dry the block. Alternatively, if glass remains, repeat the process again by
soaking the block in hydrofluoric acid for a further 20 min. (Caution: Hydrofluoric acid is a caustic chemical. Wear
gloves, and handle with care in a fume hood.)
2.4.3
Trimming the block
- Place the epoxy block on a microscope stage and obtain a bright field image (at low magnification) of the block
surface including the cell of interest.
- Using the location address on the gridded coverslip and the bright field images obtained when fluorescence images
were taken (see 2.3), check that the cell of interest is still present.
- Trim the epoxy block according to the address on the coverslip so that the block contains the cell of interest.
2.4.4
Sectioning the specimen
- Make thin slices (70-80 nm thickness) of the specimen using an ultramicrotome (Leica Biosystems, Nussloch,
Germany) with a diamond knife according to the manufacture’s protocol.
- Collect serial sections onto Formvar-coated copper slit grids. In our studies, a special 3-slit grid (No. 2486, NisshinEM Co., Tokyo, Japan) is used.
2.4.5
Observation using an electron microscope
- Incubate the sections with 4% uranyl acetate for 15-60 min to stain cell structures.
- Wash the sections with distilled water three times.
- Incubate the sections with a lead citrate solution for 1 min (a ready-to use solution is available from Sigma-Aldrich).
- Wash the sections with distilled water three times.
- Desiccate the sections in a drying chamber containing silica gel.
- Prepare a printout of a series of fluorescence 3D images and/or a bright field image before EM observation. This
printout makes it easier to find the target cells and/or subcellular structures of interest in the target cell during EM
observation.
- Observe sections using a transmission electron microscope at an appropriate voltage, and find the target
cells/structures of interest in the sections by referring to the fluorescence or bright field images on the printout.
- Obtain image data of the target cells/structures using an electron microscope with a CCD camera or on photographic
negative film. In our studies, a JEM-1200EXS transmission electron microscope (JEOL, Tachikawa, Japan) is used with
an acceleration voltage of 80 kV.
3. Results and Discussion
3.1
Live CLEM imaging of the fission yeast S. pombe
We applied “Live CLEM” to the study of the nuclear envelope in the fission yeast, S. pombe. In higher eukaryotes, the
nuclear envelope breaks down during mitosis, resulting in the intermixing of nuclear and cytoplasmic molecules. In
contrast, in the fission yeast, the nuclear envelope remains intact throughout its life cycle, including mitosis when
chromosomes segregate. However, during meiosis in S. pombe, we found that a peculiar phenomenon pertaining to the
barrier function of the nuclear envelope occurred; the cytoplasmic protein RanGAP1 entered the nucleus and nuclear
proteins became dispersed in the cytoplasm, as if the nuclear envelope had broken down. This phenomenon occurs only
during a short period of time in meiosis, about 6 min in second meiotic division. To examine whether the nuclear
envelope remained intact or not upon translocation of RanGAP1 during these few minutes in meiosis, we used Live
CLEM imaging [9].
S. pombe yeast cells expressing RanGAP1 fused with GFP and Ish1 fused with red fluorescent protein (RFP) were
induced to undergo meiosis, and living cells on gridded coverslips (CELLocate, Eppendorf in Fig. 3a; discontinued)
were observed using an Olympus oil immersion objective lens (PlanApo60/NA1.4) equipped on a DeltaVision
microscope system (Figs. 3a-d). In this particular example, instead of commercially available glass-bottom dishes,
single coverslips fixed on a hand-made coverslip holder (a 35 mm dish with a round hole smaller than the size of
coverslip in the bottom) were used. Coverslips were coated with lectin before use for immobilizing cells. The presence
of the grid lines does not affect the performance of fluorescence imaging in most dishes as well as those in the
coverslips alone. When the RanGAP1 was translocated into the nucleus during fluorescence live cell observation (see
2.2.3), the cell was fixed on the glass coverslip (Fig. 3b) and three-dimensional images of the cell were obtained (Fig.
3c). Then, the gridded coverslip was removed from the coverslip holder and subjected to embedding for EM
observation.
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Fig. 3 Live CLEM imaging of fission yeast. a, A bright field light microscopic image of living cells. The arrow
indicates a target cell for analysis in b and c. The dotted rectangle indicates the location of the target cell for
analysis in d. Bar, 15 μm. b, Live cell imaging of the target cell by FM. RanGAP1 fused to GFP and the nuclear
membrane protein Ish1 fused to mRFP can be observed simultaneously. Numbers indicate the time in minutes
after the beginning of observation. In this experiment, cells were fixed by glutaraldehyde at the 5 min time point,
when the RanGAP1-GFP translocated into the nucleus (arrows) of the cell. Bar, 10 μm. c, 3D optical sectioning
imaging of fixed cells by FM. Images were processed by deconvolution. An image of a single focal plane of the
target cell is shown. d, FM and EM images of the same field. A bright field image (BF) and a transmission
electron microscope (TEM) image are shown. Arrows indicate the target cell. Note that the target cell and some
surrounding cells remain; one nearby cell was lost during the preparation processes (asterisk). e, EM imaging.
TEM images are shown. Upper panels show whole-cell images of the cell indicated by the arrow in d. Nuc,
nucleus; NE, nuclear envelope. Lower panels, top left to bottom right, show serial section images of the area
indicated by rectangle in the upper panel. Bars, 1μm in the upper panel and 200 nm in the lower panel.
During the embedding process, the cell of interest remained attached but some cells were lost (compare left and right
panels of Fig. 3d; the cell marked by the asterisk was lost). Generally, the cells successfully observed by fluorescence
live-cell imaging tend to remain on the coated coverslips, probably because the cell attachment required for
fluorescence imaging is sufficient to keep the cell attached during sample preparation for EM analysis. The cell
observed by FM (indicated by arrows in Fig. 3d) was subjected to transmission electron microscopy as described in
Materials and Methods. Serial sections of the cell revealed an intact continuous nuclear envelope (Fig. 3e). This result
indicates that the nuclear envelope in S. pombe remains intact throughout meiosis II.
For Live CLEM of S. pombe, unlike mammalian cells, the sample holder must be resistant to acetone because the
protocol of Live CLEM of S. pombe requires acetone treatment. Specialized gridded-plastic or gridded glass-bottomed
dishes (Table 1) are suitable for this purpose. Gridded plastic-bottomed dishes (ibidi, Munchen, Germany, product
number 81161) while useful for most purposes, may not be appropriate for our method of Live CLEM because the
plastic bottom is not sturdy enough to allow accurate acquisition of 3D fluorescence images and EM images, as
described in Table 1.
3.2
CLEM imaging of the budding yeast S. cerevisiae
In another study, we used CLEM to investigate the structure of prions in the budding yeast, S. cerevisiae. The S.
cerevisiae Sup35 protein forms typical amyloid fibrils in vitro, and Sup35 fused with GFP (Sup35-GFP) forms visible
rod-shaped and spherical aggregates in the cytosol in vivo. To examine whether Sup35-GFP forms the fibrillar
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Fig. 4 CLEM imaging of budding yeast
cells. The subcellular structure showing
Sup35-GFP fluorescence was analyzed
by CLEM imaging. a, A fluorescence
and a bright field image of the target
cell. b, An EM image. c, Enlarged
image of the rectangle indicated in b.
structures in S. cerevisiae cells seen in vitro, Sup35-GFP expressing cells were fixed and subjected to CLEM imaging
[10].
Cells expressing Sup35-GFP were collected in a tube and fixed in a final concentration of 2% glutaraldehyde in 0.1
M phosphate buffer (pH 7.2) for 2 hrs at 4 °C. After four washes with the buffer, the cells were digested with
zymolyase 100T (Seikagaku Co., Tokyo, Japan) at a final concentration of 0.1 mg/ml in the buffer for 30-90 min at 30
°C. The cells were then placed on a gridded plastic-bottom dish (product number 81161, ibidi; Table 1) that was precoated with concanavalin A. Three-dimensional images (10 focal planes at 0.5 μm intervals) of a cell mounted in 0.1 M
phosphate buffer at pH 7.2 were obtained using the DeltaVision microscope system. During the embedding procedure,
however, several nearby cells were lost (data not shown). Fortunately, because budding yeast cells have large and
clearly distinguishable vacuoles, the positions, numbers, and shapes of the vacuoles served as landmarks of the cell of
interest. High-resolution EM imaging of the Sup35-GFP containing structure observed using FM of the cell of interest
showed that this aggregate of Sup35-GFP proteins contained bundled fibrillar structures (Fig. 4).
During the embedding procedure, while the cell of interest remained attached, several nearby cells were lost. Unlike
fission yeast cells, budding yeast cells attach to the dish surface at a single point because of their spherical shape.
Because of this, these cells are lost more easily than fission yeast cells.
4. Perspectives
FM enables specific molecules of interest to be visualized in living cells, while EM serves as the most appropriate
method to analyze the cellular ultrastructure. Live CLEM imaging is a powerful technique which combines the
advantages of these two forms of microscopy. The combination of FM and EM enables the molecular dynamics
obtained using FM to be correlated with the subcellular architecture identified using EM and will help elucidate aspects
of cell structure at the molecular and subcellular levels [9-11]. Fission and budding yeasts have served as ideal
experimental systems in which classical genetics, cell biology, molecular genetics and biochemistry techniques can be
applied. Such a variety of methodological approaches highlights the advantages of using yeast in biological studies. The
application of Live CLEM imaging will make yeast systems even more valuable in future biological research.
Acknowledgements We thank T. Kojidani, H. Osakada and C. Mori for their technical assistance and their efforts to successfully
implement the Live CLEM imaging system, and D. B. Alexander for critical reading of the manuscript. This work was supported by
grants from the MEXT of Japan (to H.A, Y.H. and T.H.).
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