ARTICLE
1234567890():,;
https://doi.org/10.1038/s42003-020-1095-x
OPEN
Correlation of fluorescence microscopy, electron
microscopy, and NanoSIMS stable isotope imaging
on a single tissue section
Céline Loussert-Fonta 1 ✉, Gaëlle Toullec1, Arun Aby Paraecattil2, Quentin Jeangros3, Thomas Krueger
Stephane Escrig1 & Anders Meibom 1,4
1,
Correlative light and electron microscopy allows localization of specific molecules at the
ultrastructural level in biological tissue but does not provide information about metabolic
turnover or the distribution of labile molecules, such as micronutrients. We present a method
to directly correlate (immuno)fluorescent microscopy, (immuno)TEM imaging and NanoSIMS isotopic mapping of the same tissue section, with nanometer-scale spatial precision.
The process involves chemical fixation of the tissue, cryo sectioning, thawing, and air-drying
under a thin film of polyvinyl alcohol. It permits to effectively retain labile compounds and
strongly increases NanoSIMS sensitivity for 13C-enrichment. The method is illustrated here
with correlated distribution maps of a carbonic anhydrase enzyme isotype, β-tubulin proteins,
and 13C- and 15N-labeled labile micronutrients (and their anabolic derivates) within the tissue
of a reef-building symbiotic coral. This broadly applicable workflow expands the wealth of
information that can be obtained from multi-modal, sub-cellular observation of biological
tissue.
1 Laboratory
for Biological Geochemistry, School of Architecture, Civil and Environmental Engineering, Ecole Polytechnique Fédérale de Lausanne (EPFL), CH1015 Lausanne, Switzerland. 2 Greenlight Syntax Sarl, Avenue de Morges 41, 1004 Lausanne, Switzerland. 3 Photovoltaics and Thin-Film Electronics
Laboratory, Institute of Microengineering, École Polytechnique Fédérale de Lausanne (EPFL), CH-2002 Neuchâtel, Switzerland. 4 Center for Advanced
Surface Analysis, Institute of Earth Sciences, University of Lausanne, CH-1015 Lausanne, Switzerland. ✉email: celine.loussert@epfl.ch
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
1
ARTICLE
S
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
tate-of-the-art studies of complex biological processes often
combine multiple experimental methods and employ a
variety of optical-, electron-, and ion microscopy techniques
to image tissues at the single-cell level. These techniques have
enabled advances in biology and life science by providing information about the localization and distribution of specific molecules (e.g., fluorescence in situ hybridization (FISH)1 and
immunolabelling2), as well as anabolic turnover and cellular
exchange processes (e.g. NanoSIMS3,4), together with ultrastructural information in even the most complex biological tissues. When these techniques are used in combination, the
approach is commonly referred to as correlative microscopy5.
Although correlative microscopy now features prominently in
studies of biological tissue, important limitations still exist with
regard to how well different types of imaging information can be
correlated. These limitations are primarily due to sample preparation constraints. For example, in correlative light- and electron microscopy (CLEM) most protocols involve two main steps,
starting with live or fixed-cell fluorescence imaging, followed by
sample preparation for electron microscopy (EM)6. Classical EM
sample preparation involves chemical fixation, heavy metal
staining, dehydration with solvents, resin embedding, and subsequent (ultra) thin sectioning (Fig. 1 left panel). Following this
treatment, the soluble compounds in the cell (i.e., cytosolic
components) are either thoroughly displayed7 or completely lost,
leading to a shrinkage of the tissue. Altogether it is a different
sample compared to the material that was observed previously by
fluorescence microscopy. To overcome this problem, a sample
preparation method developed by Prof. Tokuyasu8 is often used
in CLEM9,10, permitting fluorescence and EM to be carried out
on the same section9. This method involves chemical fixation
similar to classical preparation, cryo sectioning, and thawing at
room temperature. By avoiding dehydration and resin embedding, the Tokuyasu method minimizes morphological artifacts (in
particular tissue shrinkage) and chemical modifications at the
molecular level, preserving the (auto-)fluorescence properties of
the sample10 and (to variable degree) its antigenicity, thus permitting a variety of antibody-labeling.
In the last 10–15 years, ultrahigh resolution (ca. 100 nm lateral
resolution) quantitative ion microprobe imaging (NanoSIMS),
combined with experiments that introduce stable isotopic (e.g.,
13C and 15N) and/or elemental labels into a tissue, has made it
possible to study anabolic turnover and track-specific molecules
with sub-cellular resolution. NanoSIMS imaging has found
applications across numerous disciplines within the environmental, biological, and life sciences3,4,11. Nevertheless, it is still
not possible to correlate information obtained with all three
imaging techniques (i.e., fluorescence microscopy, EM, and
NanoSIMS) from one-and-the-same section of a biological tissue.
Such a capability would represent a major breakthrough in correlative microscopy12 because it would permit structural, molecular, and anabolic/metabolic information to be directly
correlated at the subcellular level. Here we present a methodology, building upon the Tokuyasu13 method, that enables direct
correlation of (immuno)fluorescent microscopy, (immuno)TEM,
and NanoSIMS ultra high-resolution stable isotopic mapping of
the same biological tissue section (Fig. 1 right panel).
We demonstrate this additional level of correlative microscopy
on tissue from a symbiotic coral (here Stylophora pistillata,
Fig. 2a). Reef-building corals are highly complex organisms that
consist of a wide range of tissue and cell types. In these organisms, the ectoderm and endoderm of both oral and aboral layers
are separated by a hydrogel-like matrix (mesoglea; black arrowheads in Fig. 2b, c). Many of the endodermal cells host symbiont,
photosynthesizing dinoflagellate algae inside symbiosomes14. A
diverse community of bacteria15 adds to the complexity of this
2
symbiotic organism, which is referred to as the “coral holobiont”.
The structural complexity of symbiotic corals thus represents a
methodological challenge. At the same time, it conveniently
provides the opportunity to demonstrate our sample preparation
and correlative microscopy approach on a range of different
matrices within a single biological tissue.
Results
The method presented here builds on the Tokuyasu method with
a number of significant improvements that results in compatibility with NanoSIMS stable isotopic imaging, while allowing
efficient immunolabeling and preservation of tissue ultrastructure
at the TEM level.
Preserving antigenicity with light chemical fixation. Some
degree of chemical fixation is required to preserve the tissue and
cell ultrastructure16. One of the commonly used fixatives for EM
is a mixture of 2.5% glutaraldehyde and 4% formaldehyde in
Sorensen buffer. With this combination, the rapidly penetrating
monoaldehyde temporally fixes the specimen until the slower
penetrating dialdehyde irreversibly crosslinks proteins.17 This
crosslinking preserves the ultrastructure of the sample but has
deleterious effects on immunolabelling, because it interferes with
the antigen epitopes (partially or totally). In order to minimize
the loss of antigenicity while preserving the capability to obtain
high quality ultrastructural imaging by TEM, we performed a
series of cryo preparations with increasing concentration of glutaraldehyde (from 0 to 2.5% (vol/vol)) and 4% formaldehyde. In
the context of our work, we found that a mix of 0.5% glutaraldehyde and 4% formaldehyde preserves the tissue ultrastructure
(TEM-imaging resolution) and optimizes the preservation of
tissue antigenicity; this fixation procedure can be adapted to
specific biological tissue and antibodies.
NanoSIMS compatibility. Following tissue fixation, the classical
Tokuyasu method13 involves cryo protection, cryo sectioning,
and air drying. The drying step requires embedding of the wet
section in a methyl cellulose uranyl acetate film (MCUA; Fig. 3a),
which prevents the section from collapsing and damage to the
ultrastructure. However, this film represents an almost impenetrable physical obstacle for NanoSIMS imaging, which requires
that the primary ion beam is capable of removing any coating and
begin sputtering secondary ions from the sample itself on a short
time scale (order of minutes). The MCUA film cannot be
removed on a time scale that renders NanoSIMS imaging feasible.
Therefore, to benefit from the advantages offered by the
Tokuyasu sample preparation method and be able to perform
NanoSIMS imaging on the same tissue section, it was necessary to
develop a technique to coat and dry a wet cryo (ultra) thin section
with an extremely thin film. This film must maintain sample
integrity and preserve ultrastructure during air drying and be
easily removed during pre-sputtering to enable efficient NanoSIMS imaging. As we show in the following, spin coating with a
polyvinyl alcohol (PVA) aqueous solution as embedding medium
achieves these objectives and has a number of added advantages.
PVA in aqueous solution has previously been used as an
alternative to methylcellulose for cryo section embedding18. PVA
is highly hydrophilic19 and thus reduces the air/water surface
tension during the drying process. It is also characterized by low
viscosity (~4 mPa s) compared to the conventional methylcellulose uranyl acetate solution (~1000 mPa s), thus allowing
formation of thin films. The conventional method for the drying
step in the Tokuyasu method makes use of a wire loop to hold the
cryo section while embedding it in a drop of MCUA. This drop is
then gradually sucked up by contact with a filter paper, as the
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
ARTICLE
Fig. 1 Comparison of the classical resin embedding workflow (left panel) and the cryo sectioning methodology introduced here (right panel). The
imaging techniques enabled by each workflow are indicated at the bottom of the figure and are performed on identical ROIs illustrated by the red squares.
Regarding cryo preparation, the correlative workflow is performed sequentially, starting with (immuno)fluorescence microscopy on wet cryo section, then
followed by (immuno)electron microscopy, and NanoSIMS imaging on the same section that is spin dried and coated with polyvinyl alcool. It has to be
noted that the cryo preparation process can be performed in 2 days, versus an entire week for the classical resin-embedding preparation.
wire loop is swept across its surface until no more MCUA
solution can be removed from the loop area. This procedure
requires substantial dexterity and produces films with a thickness
on the order of 75 nm on top of the thin section (Fig. 3a).
Introducing a low viscosity PVA aqueous solution as embedding
medium yields a thinner film on a wet cryo section when using a
spin dryer (cf. Online Methods) (Supplementary Fig. 1). With this
approach, which does not require the same level of training and
skill as the formation (by hand) of a MCUA film, a film of PVA
with a thickness of 15–20 nm can be rapidly and reproducibly
formed on top of a wet thin section (Fig. 3b). The ultrastructure
of the tissue is comparably maintained by the two drying methods
(Fig. 3c, d). Note the stark difference in the width of the gel-like
mesoglea in Fig. 3c, d compared to Fig. 2c; dehydration with
ethanol during classical sample preparation results in a strong
shrinkage of hydrated matrices, which is avoided with our (and
the Tokuyasu) method.
Correlating NanoSIMS isotopic imaging with TEM- and
fluorescent microscopy. In order to correlate (immuno)fluorescence-, (immuno)EM-, and NanoSIMS isotopic imaging, a sample holder suitable for all three imaging modalities is required.
Here we used TEM grids as carriers of the cryo thin sections. A
wet, thin section supported by a TEM grid is fully compatible
with fluorescent microscopy10. However, the fragility of TEM
grids means that they require a protective support during spin
drying. We used a polytetrafluoroethylene (PTFE)-coated rubber
septa with a hydrophilic top-surface that maintained the TEM
grid in place during spin coating (Supplementary Fig. 1); following spin drying the TEM grid was easily handled for subsequent TEM and NanoSIMS-imaging technics (Fig. 1).
With this workflow, the same cryo thin section (with a
thickness of ca. 100 nm) is sequentially imaged in wet conditions
by fluorescence microscopy, and then, after drying, by TEM and
NanoSIMS. This is demonstrated in Fig. 4, in which—for
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
3
ARTICLE
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
Fig. 2 The symbiotic coral Stylophora pistillata used as model organism in this study. a A small coral colony. b Histological section of a decalcified piece
of coenosarc tissue (i.e., between two polyps). The coenosarc is divided into the oral and aboral tissues, subdivided into oral ectoderm, oral endoderm,
aboral endoderm, and aboral ectoderm (calicoderm). The oral ectoderm is directly facing seawater and the calicoderm is facing the skeleton. Oral
ectoderm/oral endoderm and aboral endoderm/calicoderm are separated by a gel-like mesoglea (black arrows). Many oral endodermal cells host
photosynthesizing dinoflagellate algae symbionts (Symbiodinium; one marked by an asterisk) surrounded by a symbiosome membrane. c The mesoglea
interface between the oral ectoderm and oral endoderm is shown in TEM following classical sample preparation (cf. Online Methods). Note the narrow
width of the mesoglea. White arrowhead: nucleus; asterisk: dinoflagellate symbiont; black arrowhead: mesoglea; OEct oral ectoderm, OEnd oral endoderm,
Col coelenteron, Ab aboral endoderm. Cal calicoblastic ectoderm. Scale bars: a 1 cm; b 10 μm; and c: 5 μm.
Fig. 3 Secondary electron scanning electron micrographs of focused ion beam-prepared (FIB) cross-sections of the cryo sections deposited on formvar
carbon films (supported on Cu grids) and coated with either MCUA (a) or spin-coated with PVA (b). Both thin sections were stained with heavy metals
to enhance image contrast. To avoid charging and facilitate contrast differentiation with the overlying protective layer of carbon deposited in the FIB
section, a 10 nm layer of Au was deposited on top of the MCUA or PVA films. Gray scale intensity profiles taken at the positions indicated by red arrows in
a and b show the difference in thickness of MCUA and PVA films (indicated as shaded areas). c and d TEM images of the oral ectoderm–endoderm
interface in S. pistillata obtained with the MCUA layer deposited with normal Tokuyasu sample preparation and with the thin PVA film (this method),
respectively. Note the width of the mesoglea (black arrowheads). White arrowheads indicate nuclei; asterisks: symbionts; OEct oral ectoderm, OEnd oral
endoderm. Scale bars: a and b: 100 nm; c and d: 5 μm.
4
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
ARTICLE
Fig. 4 The correlative workflow combining immuno-fluorescence microscopy and immuno-electron microscopy with NanoSIMS imaging on the same
tissue section of a coral after incubation with H13CO3− and 15NO3− (and therefore enriched in 13C and 15N); cf. Online Methods. The cryo section (ca.
100 nm thick) was imaged after simultaneous immunolocalization of carbonic anhydrase with anti-Alexa-associated secondary antibodies (a and b) and βtubulin observed in TEM by 10 nm gold particles associated to the secondary antibody (c and d). The fluorescence microscope images (a and b; zoom in a)
and the TEM micrographs (c and d; zoom in c) exhibit identical areas of the same thin section. e and f are NanoSIMS images showing the 13C and 15N
distributions in the interior of a dinoflagellate symbiont. The area imaged is indicated by a red square in c. Scale bars: a and c: 5 μm; b and d: 500 nm; e and
f: 2 μm.
illustration purposes—the localization of one isotype of carbonic
anhydrase (CA) enzyme is revealed by fluorescence microscopy
(Fig. 4a, b), the localization of the β-tubulin proteins by TEM
(Fig. 4c, d), and the distribution of 13C- and 15N-enrichments by
NanoSIMS imaging (Fig. 4e, f) on the exact same area of a thin
section of the symbiotic coral S. pistillata, incubated for 6 h in
seawater with 2 mM H13CO3− and 3 μM 15NO3− under natural
daylight illumination (cf. Online Methods).
CA is a metallo-enzyme essential to photosynthesis in the
symbiont dinoflagellate algae, where it reversibly catalyzes the
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
5
ARTICLE
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
conversion of CO2 into carbonic acid, bi-carbonate ions, and
protons. The anti-CA2 antibody used for this experiment was
raised against a peptide sequence from the human CA2 protein.
We matched several proteins from S. pistillata and Cladocopium (previous nomenclature Symbiodinium clade C)20 by
BLASTp. A maximal identity score of 50.2% was obtained for
the CA2 isoform X1 (accession number XP 022794253.1) for
the host coral, and of 27.2% for the CA14 isoform associated
with the dinoflagellate symbionts (accession number
OLQ02879.1). With this anti-CA2 antibody we observed CA
to be concentrated in the pyrenoid structure and in secondary
starch granules inside dinoflagellate cells, and at the vicinity of
the symbiosome membranes (Fig. 4a, b); we did not observe
distinct localization of CA in the coral host tissue, most likely
due to the non-denaturizing conditions used for the immunolabelling. Negative control samples exhibited no fluorescence.
After spin drying and deposition of a ca. 20 nm film of PVA
on the thin section, the same region of the section was
subsequently imaged by TEM to reveal the location of βtubulin, using a secondary antibody linked to 10 nm gold
particles. The commercial anti-β-tubulin antibody used for this
work is known to react with a broad spectrum of β-tubulin
isotypes21, many of which are present in both S. pistillata and in
Symbiodiniaceae. We identified three sites of β-tubulin
localization in the coral holobiont: the motile cilia on the outer
surface of the host’s oral ectoderm (i.e., the coral host;
Supplementary Fig. 2A and A’) and in the symbionts, inside
the pyrenoids, in a shell-like structure embedded in the primary
starch sheet and surrounding the central pyrenoid matrix
(Fig. 4c, d; Supplementary Fig. 2B), and around starch granules
(Supplementary Fig. 2C). We speculate that the structures
inside the pyrenoid containing β-tubulin proteins might
contain pyrenoid tubules22. The co-localization of CA and βtubulin proteins in a layer surrounding the electron dense
pyrenoid matrix is consistent with their presumed role in
concentration and delivering of CO2 to the Rubisco-rich center
of the pyrenoid, as also described for the marine diatom
Phaeodactylum tricornutum23.
Finally, the same thin section was coated with ca. 10 nm Au
(standard procedure to avoid surface charging) and transferred
to the NanoSIMS ion-microprobe in which the distribution of
13C- and 15N-enrichments were mapped in the same region24.
Following pre-sputtering that rapidly removed the Au coating
and the PVA film, the primary Cs+ ion beam was focused to
about 100 nm and rastered across the sample surface producing
of secondary ions, notably 12C12C−, 13C12C−, 12C15N−, and
12C14N−. These ions were extracted, separated from potentially
interfering ions in the mass spectrometer, and individually counted
in electron-multiplier detectors3 (cf. Online Methods). This
permitted 13C- and 15N-enrichments to be quantified through the
count-rate ratios 13C12C−/12C12C− and 12C15N−/12C14N−, respectively. Isotopic enrichments are presented in the δ-notation, i.e., in
parts-per-thousand relative to a control sample of normal isotopic
composition, prepared and analyzed in an identical manner.
The lateral resolution of a NanoSIMS image is sufficient to
allow precise correlation with both the TEM and the fluorescent
microscopy images (Fig. 4; Supplementary Fig. 3). NanoSIMS
images revealed enrichment levels of 13C and 15N inside the
symbiont nucleus (except in the condensed chromosomes) and
cytoplasm (Fig. 4; Supplementary Fig. 3). The chloroplasts and
the shell surrounding the pyrenoid matrix (which also contained
CA and β-tubulin proteins) were enriched in 13C, consistent with
an active role in C-metabolism of these organelles25. The
pyrenoid matrix exhibited both 13C and 15N enrichments
(Supplementary Fig. 3). These observations are consistent with
6
the notion of a high concentration of constantly renewing
RubisCO proteins located in this matrix22,26.
NanoSIMS imaging—advantages in comparison with conventional sample preparation. Our sample preparation method
creates several important consequences for NanoSIMS imaging.
Because our method avoids ethanol dehydration and resin
embedding, the chemical composition of the sample (i.e., the
matrix) is very different from a resin-embedded section prepared
with classical protocols. We observed that the ionization efficiency, and hence the count rate of key ions such as CN−, is
higher from the matrix created with our method. Supplementary
Fig. 4 shows a qualitative comparison between the 12C14N−
count-rates obtained with identical analysis conditions on two
comparable coral tissue sections (i.e., similar sample areas, tissue
regions, and symbiont density), prepared classically vs. our
method (Fig. 1). With the sample matrix generated with our
method, the 12C14N− count-rate is about 1.8 times higher than
with classical sample preparation, permitting to either obtain
better counting statistics with the same analysis time, or reduce
the analysis time correspondingly.
Furthermore, in a direct comparison between coral tissue
sections prepared classically (i.e., with dehydration and resin
embedding) and with our method (i.e., without dehydration and
resin embedding) we measured up to about a factor of 10 higher
13C-enrichments in the latter (Fig. 5). Dehydration (e.g., with
ethanol) contributes to a wash-out of labile molecules not
immobilized during the chemical fixation step (such as e.g.
bicarbonate ions). Moreover, resin embedding introduces a large
amount of C with normal isotopic composition and dilutes tissue
13C-enrichments3,27. This resulted in a strong discrepancy
between the 13C-enrichments originally present in a given tissue
and those measured with the NanoSIMS on a classically prepared
tissue section. With the method introduced here, labile C-rich
compounds were not lost to the same degree from the sample and
the absence of resin embedding avoided further dilution of the
remaining 13C-enrichments.
The loss and dilution of the 13C-enrichment with dehydration
and resin embedding depended on tissue/organelle structure and
density. The biggest difference in measured 13C-enrichments
between tissue prepared classically vs. our method was observed
in the gel-like mesoglea and in coral host cells (Fig. 5a, b). In the
dinoflagellate algae the effect was less pronounced because their
13C-enrichments were primarily concentrated in starch grains
and lipid droplets28, which were less affected by dehydration and
resin penetration. Note that, because resin contains little nitrogen,
its effect on 15N/14N ratios was insignificant (Supplementary
Fig. 5).
Short of a pure cryo sample preparation chain (i.e., starting
with high-pressure freezing and keeping the sample frozen
through all subsequent preparation steps) that would preserve
all cellular components in place, the sample preparation
method presented here (Fig. 1b) preserves labile components
in situ to the highest possible degree. These labile components
are often essential metabolites and/or precursors for molecules
with higher structural order produced in anabolic processes29.
However, it takes time for anabolic processes (typically minutes
to hours), to transfer a measurable (with the NanoSIMS)
isotopic enrichment from a labile precursor molecule into
structural molecules30,31. Therefore, with experiments in which
labile components enter the tissue but are not allowed time to
be converted by anabolic processes, our preparation method
provides the best chance of imaging the distribution of
isotopically enriched labile compounds, at least at the tissue
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
ARTICLE
Fig. 5 Quantitative NanoSIMS analyses of 13C-enrichments in two
different coral colonies incubated with H13CO3− under identical
experimental conditions (cf. Online Methods). Comparison between
classical sample preparation (i.e., with resin embedding) and the cryo
sample preparation protocol developed in this study was made for (a) coral
host cells, (b) mesoglea, and (c) dinoflagellate symbionts (chemical fixation
was 2.5% glutaraldehyde and 4% formaldehyde in both preparations). This
sample preparation protocol systematically preserved stronger 13Cenrichments in the tissue, by up to a factor of 10, except in the
dinoflagellate symbionts, where 13C-enrichments were primarily localized in
starch granules and lipid droplets, which were less affected by resin
dilution. The dashed lines indicate the detection limit for the NanoSIMS,
defined here as three standard deviations obtained by analyzing similar
regions in unlabeled control tissue (n = 9). The statistical differences
between the treatments were calculated by one-way ANOVA. The
annotation of p-value significance level is: “***”[0, 0.001]; “**”(0.001,
0.01]; “*”(0.01, 0.05]; “.“(0.5, 0.1]; “” (0.1, 1]; ns non-significant.
Discussion
We have presented a sample preparation method that allows
direct correlation of (immuno)fluorescent microscopy, (immuno)
TEM, and NanoSIMS isotopic imaging of a single tissue thin
section. We have demonstrated this capability on reef-building
symbiotic coral tissue, which represents a methodological challenge because it contains a broad range of biological matrices,
from hydro-gel structures to dense starch granules and lipid
droplets. Our method creates a number of advantages for
NanoSIMS isotopic imaging, notably enhanced ionization efficiency of key secondary ions, and the possibility to map the
distribution of isotopically labeled labile components, which are
partly retained with our method but lost during classical sample
preparation. Our method is applicable to a wide range of biological tissue. This additional level of sophistication in correlated
microscopy will find application across the life- and biological
sciences, as well as in many branches of the environmental
sciences.
Methods
Coral maintenance and experimental design. Specimens of S. pistillata were
collected from the shallow waters (4–8 m) of the Gulf of Aqaba in Eilat, Israel under
permit 2019/42143 of the Israel Nature and Parks Authority. Experiments were conducted in the outdoor Red Sea Simulator system at the Interuniversity Institute (IUI)
for Marine Sciences with corals maintained in air-bubbled aquaria (2 L) at ambient
water temperature (28 °C) and reduced solar irradiance (ca. 200 µmol m−2 s−1 at
midday under canopies). Three different mother colonies were selected according the
similarity of their sizes. For the labeling pulse, two corals were incubated in seawater
spiked with 2 mM NaH13CO3 98 at.% (Sigma-Aldrich, Switzerland) and 3 µM K15NO3
98 at.% (Sigma-Aldrich, Switzerland)32. A piece of an unlabeled coral colony was
used as reference to assess the natural isotopic ratio in the tissue of interest. This coral
was incubated in natural sea water under similar illumination conditions. The
experiments were conducted between 10 h 30–16 h 30. The temperature and light
illumination were recorded each hour during the 6 h. At the end of the isotopic pulse,
fragments of branches at half centimeter from the apical tips were removed, broken
into pieces ~1 cm in linear dimension, and immediately transferred into phosphate
buffer (0.1 M pH 7.4) with 9% sucrose, containing 4% formaldehyde and glutaraldehyde in different concentrations: 0%, 0.5%, 1%, and 2.5%. Fixation was done
at room temperature for 2 h after which the samples were stored in fresh fixative
overnight at 4 °C.
level. Conversely, anabolic tissue turnover can be studied by
pulse-chase experiments, in which isotopically labeled, labile
precursor molecules are permitted to enter the tissue (pulse)
and be either metabolized or flushed out of the tissue again (the
chase) prior to NanoSIMS imaging.
Classical tissue preparation. After fixation, the samples were decalcified in 0.5 M
EDTA solubilized in 0.1 M Sorensen buffer (pH 7.4) containing 0.4% formaldehyde
at room temperature for 4 days. Coenosarc tissues were then cut into 1 mm2 pieces
and post stained in 2% osmium aqueous solution for one hour in the dark. After
rinse in water, samples were dehydrated in a series of ethanol concentrations,
ranging from 10% to 100% then infiltrated with SPURR resin (Electron Microscopy
Sciences, USA) before polymerization at 60 °C for 24 h. Thin sections of 90 nm
were cut and mounted onto a Formvar film-coated, carbon-stabilized 100 mesh
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
7
ARTICLE
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
copper finder grid (Electron Microscopy Sciences, USA) or 200 nm thin sections on
silicon wafers (Siltronix, Archamps, France) (Supplementary Fig. 1).
Cryo preparation. After fixation and decalcification, coenosarc tissues were casted
in 12% gelatin (which does not penetrate into the sample) in 0.1 M Sorensen buffer
pH 7.4 and cut into 1 mm3 cubes. Then each sample was cryo protected by
infiltration with 2.3 M sucrose (a non-permeating cryoprotectant) in 0.1 M Sorensen buffer pH 7.4 overnight at 4 °C; in this condition the samples can be stored
for months. Before cryo thin sectioning, sucrose infiltrated and gelatin-embedded
coenosarc pieces were oriented and mounted on aluminum pins under a binocular
and immediately plunged into liquid nitrogen (Fig. 1d).
Cryo sectioning, immunolabelling, and microscopy. Pins with frozen tissue were
mounted in a cryo ultramicrotome (Ultracut UC6/FC6, Leica Microsystems, Austria).
Coenosarc blocks were trimmed at −90 °C with a Cryotrim diamond knife (Diatome,
Switzerland) and sections of 100 or 200 nm thick were cut at −90 °C with an immuno
diamond knife (Diatome, Switzerland). These sections were picked up with a drop of
a mixture containing an equal volume of 2% methylcellulose and 2.3 M sucrose33 to
minimize material extraction (compared to pick-up in pure sucrose), warmed up to
room temperature, and transferred onto a Formvar film-coated, carbon-stabilized 100
mesh copper finder grid or silicon wafers.
Histological imaging. Both resin and cryo sections were mounted onto Superfrost
Adhesion microscope slides (ThermoFisher, USA) and stained with a mixture of 1% s
toluidine blue and 1% basic fuchsin in water34 30 s on 60 °C heat plate. After rinsing
in distillated water, the sections were imaged with an upright microscope DM 5500
equipped with CCD camera DFC 3000 B/W (Leica Microsystems, Switzerland)
controlled with the Leica software LAS-X (Leica Microsystems, Switzerland).
Beta-tubulin immuno-gold labeling. Cryo thin sections collected onto TEM grids
were incubated 30 min on gelatin 2% in 0.1 M Sorensen buffer, pH 7.4, at 37 °C to
remove the 12% gelatin (used for the embedding of the sample) from the section, then
incubated 5 min in PBS buffer containing 1% BSA (Aurion, The Netherlands) as a
blocking step to avoid nonspecific labeling. Sections were floated for 1 h on a drop of
mouse anti-beta-tubulin antibody (Sigma-Aldrich, Switzerland) diluted at 1/50 in PBS
containing 1% BSA. After washing with 0.1% BSA in PBS, the samples were incubated
for 1 h with 10 nm colloidal gold-conjugated secondary antibodies, goat anti-mouse
(Aurion, The Netherlands) diluted 1:30 in 1% BSA/PBS. The ultrathin cryo sections
were then fixed in 1% glutaraldehyde in PBS for 10 min to further stabilize them,
followed by eight washes in distilled water, 2 min each. Finally, samples were dried
and PVA embedding was done as will be described in the subsection “Drying and
embedding of the cryo sections for NanoSIMS analysis”.
CA immuno-fluorescence labeling. Cryo thin sections were processed as described
in the previous subsection with a rabbit anti-CA (reference 100-4157, Rockland, USA)
diluted 1/50 as the first antibody and a donkey anti-rabbit linked with Alexa 568
(ThermoFisher, USA) as the secondary antibody. To avoid any autofluorescence, the
sections were not post-fixed before imaging. They were then were mounted between a
Superfrost Adhesion microscope slides (ThermoFisher, USA) and a 18 × 18 mm
Corning® cover glass (Merk, Germany) with PBS containing DAPI10 as mounting
medium. The glass slide/coverslip chambers were sealed with commercial nail polish.
Prior to imaging, the samples were kept in the dark, inside a humidity chamber at
4 °C for a maximum of 6 h. Imaging was done using Zeiss LSM 700 inverted confocal
microscope (Zeiss, Germany) equipped with Axiocam MRm (B/W) camera (Zeiss,
Germany) and operated with the software Zen 2009 (Zeiss, Germany). After imaging,
the glass slide/coverslip chambers were disassembled.
When double immunolabelling was performed, the sections were floated on a
drop of solution containing both primary antibodies. The secondary antibodies were
also mixed in the second incubation step. All other incubation steps were performed
as described previously. When immunolabelling was performed on cryo sections
mounted on silicon wafers, glass bottom 24-well-imaging plates (MatTek, USA) were
used during epifluorescence imaging. The silicon wafers were placed upside down
inside the plate chambers onto a 20 µl drop of PBS containing DAPI. Before imaging,
the samples were kept in the dark, inside a humidity chamber at 4 °C for a maximum
of 6 h. Imaging was done using Zeiss LSM 700 inverted confocal microscope (Zeiss,
Germany) equipped with Axiocam MRm (B/W) camera (Zeiss, Germany) and
operated with the software Zen 2009 (Zeiss, Germany). After imaging, samples were
rinsed in water prior to drying and PVA embedding, as described next.
Drying and embedding of the cryo sections for NanoSIMS analysis. After
rinsing in Sorensen buffer (pH 7.4) the antigen/antibody bonds were stabilized in
1% glutaraldehyde in PBS for 10 min. After eight washes in distilled water (2 min
each), the sections were incubated for 5 min in aqueous solution of 3% PVA
(Sigma-Aldrich, Switzerland) followed by spin drying at 66 rotations per second for
45 s to produce a thin film of PVA on the wet section (Supplementary Fig. 1). If
silicon wafers or coverslips were used as support for the cryo thin sections during
spin drying, no special sample holder was required. When TEM grids were used as
support, these had to be protected. A PTFE-coated rubber septa (Agilent, USA) was
8
found to be an ideal support for TEM grids, because the PTFE side is providing a
hydrophilic top-surface on which a grid was kept in place during spin drying,
following which it was easily removed for subsequent imaging in TEM and
NanoSIMS (Supplementary Fig. 1).
SEM imaging. Secondary electron SEM images of FIB-prepared cross sections
(Fig. 3a, b) were obtained in a Zeiss NVision 40 (Zeiss, Germany). The procedure
for cross-sectional imaging involved the deposition of a carbon protection layer,
first with the electron beam and then with a gallium beam, on Au-coated layer
stacks. Cross-sections were prepared with a gallium ion beam current of 3–1.5 nA
(at an acceleration voltage of 30 kV). In-lens secondary electron SEM images were
acquired with a beam voltage of 2 kV.
TEM imaging. Thin sections were imaged with a Tecnai-12 transmission electron
microscope (ThermoFisher, USA) operating at 100 kV with a FEI eagle camera
(ThermoFisher, USA) using TIA software (ThermoFisher, USA).
NanoSIMS imaging. Thin sections were gold-coated to a thickness of ca. 10 nm
prior to mapping of 13C- and 15N-enrichments by a NanoSIMS 50 L ion
microprobe (Cameca, France) using a 16 keV primary Cs+ ion beam. Following
pre-sputtering, the primary beam was focused to a spot-size of around 100 nm and
rastered across the sample surface in selected regions (guided by fluorescence and
TEM imaging) with areas of 40 × 40 or 25 × 25 μm2, 256 × 256 pixels, and a pixel
dwell-time of 5000 µs. The secondary ions 12C2−, 13C12C−, 12C14N−, and 12C15N–
were separated from potential interferences at a mass resolution of around 9000
(Cameca definition) and counted individually and simultaneously in electron
multipliers. Each finished image consisted of five rastered images (except for the
images displayed in Fig. 4, which consist of 30 layers) added together following
drift correction using the L’Image© software developed by Dr. Larry Nittler.
Statistics and reproducibility. The NanoSIMS data presented in this work
represent 171 dinoflagellate symbionts (with an apparent diameter larger than
5 µm to ensure a close to equatorial cut through the symbiont) from two colonies,
and 26 image frames of 40 × 40 μm for host tissue and mesoglea. To assess the
effect of sample preparation on NanoSIMS measurements, we analyzed the data by
one-way ANOVA.
Reporting summary. Further information on research design is available in
the Nature Research Reporting Summary linked to this article.
Data availability
The datasets generated during and/or analyzed during the current study are available
from the corresponding author on reasonable request. Source data are avaialable as
Supplementary Data 1.
Received: 9 April 2020; Accepted: 19 June 2020;
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
Huber, D., Voith von Voithenberg, L. & Kaigala, G. V. Fluorescence in situ
hybridization (FISH): history, limitations and what to expect from micro-scale
FISH? Micro Nano Eng 1, 15–24 (2018).
de Matos, L. L., Trufelli, D. C., de Matos, M. G. L. & da Silva Pinhal, M. A.
Immunohistochemistry as an important tool in biomarkers detection and
clinical practice. Biomark Insights 5, 9–20 (2010).
Hoppe, P., Cohen, S. & Meibom, A. NanoSIMS: technical aspects and
applications in cosmochemistry and biological geochemistry. Geostand
Geoanal Res 37, 111–154 (2013).
Nuñez, J., Renslow, R., Cliff, J. B. & Anderton, C. R. NanoSIMS for biological
applications: current practices and analyses. Biointerphases 13, 03B301 (2018).
Ando, T. et al. The 2018 correlative microscopy techniques roadmap. J. Phys.
D 51, 443001 (2018).
Hauser, M. et al. Correlative super-resolution microscopy: new dimensions
and new opportunities. Chem. Rev. 117, 7428–7456 (2017).
Huebinger, J., Spindler, J., Holl, K. J. & Koos, B. Quantification of protein
mobility and associated reshuffling of cytoplasm during chemical fixation. Sci.
Rep. 8, 17756 (2018).
Tokuyasu, K. T. Immunochemistry on ultrathin frozen sections. Histochem. J.
12, 381–403 (1980).
Oorschot, V. M. J., Sztal, T. E., Bryson-Richardson, R. J. & Ramm, G. Immuno
correlative light and electron microscopy on tokuyasu cryosections. In
Methods in Cell Biology, Vol. 124, 241–258 (Elsevier, edited by Thomas
Müller-Reichert and Paul Verkade, 2014).
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-020-1095-x
10. Loussert Fonta, C. et al. Analysis of acute brain slices by electron microscopy:
a correlative light-electron microscopy workflow based on Tokuyasu cryosectioning. J. Struct. Biol. 189, 53–61 (2015).
11. Gyngard, F. & Steinhauser, M. L. Biological explorations with nanoscale
secondary ion mass spectrometry. J. Anal. At. Spectrom. 34, 1534–1545 (2019).
12. Decelle, J. et al. Subcellular chemical imaging: new avenues in cell biology.
Trends Cell Biol. S0962892419302211 (2020) https://doi.org/10.1016/j.
tcb.2019.12.007.
13. Tokuyasu, K. T. A technique for ultracryotomy of cell suspensions and tissues.
J. Cell Biol. 57, 551–565 (1973).
14. Wakefield, T. S. & Kempf, S. C. Development of host- and symbiont-specific
monoclonal antibodies and confirmation of the origin of the symbiosome
membrane in a Cnidarian–Dinoflagellate symbiosis. Biol. Bull. 200, 127–143
(2001).
15. Rosenberg, E., Koren, O., Reshef, L., Efrony, R. & Zilber-Rosenberg, I. The role
of microorganisms in coral health, disease and evolution. Nat. Rev. Microbiol.
5, 355–362 (2007).
16. Li, Y. et al. The effects of chemical fixation on the cellular nanostructure. Exp.
Cell Res. 358, 253–259 (2017).
17. Hayat, M. A. Principles and Techniques of Electron Microscopy: Biological
Applications (Cambridge University Press, 2000).
18. Tokuyasu, K. T. Use of poly(vinylpyrrolidone) and poly(vinyl alcohol) for
cryoultramicrotomy. Histochem. J. 21, 163–171 (1989).
19. Yang, E., Qin, X. & Wang, S. Electrospun crosslinked polyvinyl alcohol
membrane. Mater. Lett. 62, 3555–3557 (2008).
20. LaJeunesse, T. C. et al. Systematic revision of Symbiodiniaceae highlights the
antiquity and diversity of coral endosymbionts. Curr. Biol. 28, 2570–2580.e6
(2018).
21. Yang, H., Cabral, F. & Bhattacharya, R. Tubulin isotype specificity and
identification of the epitope for antibody Tub 2.1. Protein Eng. Design Sel. 22,
625–629 (2009).
22. Meyer, M. T., Whittaker, C. & Griffiths, H. The algal pyrenoid: key
unanswered questions. J. Exp. Bot. 68, 3739–3749 (2017).
23. Kikutani, S. et al. Thylakoid luminal θ-carbonic anhydrase critical for growth
and photosynthesis in the marine diatom Phaeodactylum tricornutum. Proc.
Natl. Acad. Sci. USA 113, 9828–9833 (2016).
24. Kopp, C., Domart-Coulon, I., Barthelemy, D. & Meibom, A. Nutritional input
from dinoflagellate symbionts in reef-building corals is minimal during
planula larval life stage. Sci. Adv. 2, e1500681–e1500681 (2016).
25. Roth, M. S. The engine of the reef: photobiology of the coral–algal symbiosis.
Front. Microbiol. 5, 1-22 (2014).
26. Hirel, B. & Gallais, A. Rubisco synthesis, turnover and degradation: some new
thoughts on an old problem: commentary. New Phytol. 169, 445–448 (2006).
27. Musat, N., Foster, R., Vagner, T., Adam, B. & Kuypers, M. M. M. Detecting
metabolic activities in single cells, with emphasis on nanoSIMS. FEMS
Microbiol. Rev. 36, 486–511 (2012).
28. Kopp, C. et al. Highly dynamic cellular-level response of symbiotic coral to a
sudden increase in environmental nitrogen. mBio 4, (2013).
29. Lu, W. et al. Metabolite measurement: pitfalls to avoid and practices to follow.
Annu. Rev. Biochem. 86, 277–304 (2017).
30. Gibbin, E., Banc-Prandi, G., Fine, M., Comment, A. & Meibom, A. A method
to disentangle and quantify host anabolic turnover in photosymbiotic
holobionts with subcellular resolution. Commun. Biol. 3, 14 (2020).
31. Takado, Y. et al. Imaging the time-integrated cerebral metabolic activity with
subcellular resolution through nanometer-scale detection of biosynthetic
products deriving from 13C-glucose. J. Chem. Neuroanat. 69, 7–12 (2015).
ARTICLE
32. Krueger, T., Bodin, J., Horwitz, N. et al. Temperature and feeding induce
tissue level changes in autotrophic and heterotrophic nutrient allocation in the
coral symbiosis – A NanoSIMS study. Sci Rep 8, 12710 (2018). https://doi.org/
10.1038/s41598-018-31094-1.
33. Liou, W., Geuze, H. J., Geelen, Math., J. H. & Slot, J. W. The autophagic and
endocytic pathways converge at the nascent autophagic vacuoles. J. Cell Biol.
136, 61–70 (1997).
34. Aoki, A. & Gutierrez, L. S. A simple toluidine blue—basic fuchsin stain for
spermatozoa in epoxy sections. Stain Technol. https://doi.org/10.3109/
10520296709115030 (2009).
Acknowledgements
This work was funded by Swiss National Science Foundation Grant 200021_179092 to A.
M. We thank Prof. Maoz Fine for access to his Red Sea Simulator aquarium system at the
InterUniversity Institute for Marine Science in Eilat and for insightful technical discussions. Drs. Prof. Isabelle Domart-Coulon, Savary Romain, and Nils Rädecker are
thanked for constructive criticism of earlier versions of this manuscript.
Author contributions
C.L.-F., G.T., T.K., and A.M. designed the experiments. C.L.-F. and G.T. conducted the
experiments. A.A.P. provided support for film coating. Q.J. performed FIB-SEM imaging.
S.E. provided support for NanoSIMS anaysis. C.L.-F. analyzed the samples and performed data analyis. C.L.-F. and A.M. produced the manuscript.
Competing interests
The authors declare no competing interests.
Additional information
Supplementary information is available for this paper at https://doi.org/10.1038/s42003020-1095-x.
Correspondence and requests for materials should be addressed to C.L.-F.
Reprints and permission information is available at http://www.nature.com/reprints
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in
published maps and institutional affiliations.
Open Access This article is licensed under a Creative Commons
Attribution 4.0 International License, which permits use, sharing,
adaptation, distribution and reproduction in any medium or format, as long as you give
appropriate credit to the original author(s) and the source, provide a link to the Creative
Commons license, and indicate if changes were made. The images or other third party
material in this article are included in the article’s Creative Commons license, unless
indicated otherwise in a credit line to the material. If material is not included in the
article’s Creative Commons license and your intended use is not permitted by statutory
regulation or exceeds the permitted use, you will need to obtain permission directly from
the copyright holder. To view a copy of this license, visit http://creativecommons.org/
licenses/by/4.0/.
© The Author(s) 2020
COMMUNICATIONS BIOLOGY | (2020)3:362 | https://doi.org/10.1038/s42003-020-1095-x | www.nature.com/commsbio
9