Architecture and host interface of environmental chlamydiae
revealed by electron cryotomography
Martin Pilhofer
1,2,†
,
Karin Aistleitner
3,†
,
Mark S. Ladinsky
1
,
Lena König
3
,
Matthias Horn
3,*
,
and
Grant J. Jensen
1,2,**
1
Division of Biology, California Institute of Technology, Pasadena, CA 91125, USA
2
Howard Hughes Medical Institute, Pasadena, CA 91125, USA
3
Division of Microbial Ecology, University of Vienna, Vienna, A-1090 Austria
Summary
Chlamydiae comprise important pathogenic and symbiotic bacteria that alternate between
morphologically and physiologically different life stages during their developmental cycle. Using
electron cryotomography, we characterize the ultrastructure of the developmental stages of three
environmental chlamydiae:
Parachlamydia acanthamoebae
,
Protochlamydia amoebophila
and
Simkania negevensis
. We show that chemical fixation and dehydration alter the cell shape of
Parachlamydia
and that the crescent body is not a developmental stage, but an artefact of
conventional electron microscopy. We further reveal type III secretion systems of environmental
chlamydiae at macromolecular resolution and find support for a chlamydial needle-tip protein.
Imaging bacteria inside their host cells by cryotomography for the first time, we observe marked
differences in inclusion morphology and development as well as host organelle recruitment
between the three chlamydial organisms, with
Simkania
inclusions being tightly enveloped by the
host endoplasmic reticulum. The study demonstrates the power of electron cryotomography to
reveal structural details of bacteria–host interactions that are not accessible using traditional
methods.
Introduction
All chlamydiae share an obligate intracellular life style and depend on a eukaryotic host for
replication (
Horn, 2008
). Chlamydial ancestors adapted to this life inside a host more than
700 million years ago probably thriving in ancient protists (
Horn and Wagner, 2004
;
Horn
et
al
., 2004
;
Kamneva
et al
., 2012
;
Subtil
et al
., 2013
). For a long time
Chlamydiae
were
thought to consist of only human and certain animal pathogens (the
Chlamydiaceae
). In the
past two decades, a novel class of ‘environmental’ chlamydiae have been identified in
contaminated cell cultures, in an aborted bovine fetus, in fish gills, and as symbionts of
arthropods and amoebae (
Rourke
et al
., 1984
;
Michel
et al
., 1994
;
Kahane and Friedman,
*
For correspondence. horn@microbial-ecology.net; Tel. (+43) 1 4277 76608; Fax (+43) 1 4277 876601.
**
jensen@caltech.edu; Tel.
(+1) 626 395 8827; Fax (+1) 395 5730.
†
Contributed equally.
Supporting information
Additional Supporting Information may be found in the online version of this article at the publisher’s web-site.
Published as:
Environ Microbiol
. 2014 February ; 16(2): 417–429.
HHMI Author Manuscript
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1995
;
Amann
et al
., 1997
;
Rurangirwa
et al
., 1999
;
Fritsche
et al
., 2000
;
Horn
et al
., 2000
;
Draghi
et al
., 2004
;
Kostanjsek
et al
., 2004
;
Thomas
et al
., 2006
;
Karlsen
et al
., 2008
). While
pathogenic chlamydiae are a homogeneous phylogenetic group, the genomes of
environmental chlamydiae are more diverse (
Bertelli
et al
., 2010
;
Collingro
et al
., 2011
). It is
still unclear to what extent this genomic variation manifests as variations in cell structure.
The biphasic chlamydial life cycle starts with the infection of a host cell by the elementary
body (EB) (
Horn, 2008
). After uptake by the host, the EB resides inside a host-derived
vacuole (termed ‘inclusion’) (
Hackstadt
et al
., 1997
) and differentiates into a reticulate body
(RB), the replicative developmental stage. The RB then divides several times by binary
fission before redifferentiating into EBs, which leave the host cell by lysis or exocytosis to
start a new round of infection (
Abdelrahman and Belland, 2005
;
Hybiske and Stephens,
2007
;
Horn, 2008
). An additional infectious developmental stage, the sickle-shaped crescent
body, was reported for a number of environmental chlamydiae (
Greub and Raoult, 2002
;
Lamoth and Greub, 2010
;
Nakamura
et al
., 2010
).
Once inside their host, chlamydiae perturb the organelle organization of the host cell in
various ways.
Chlamydia trachomatis
inclusions, for instance, cause fragmentation of the
Golgi (
Heuer
et al
., 2009
), which facilitates the acquisition of cholesterol and sphingomyelin
(
Hackstadt
et al
., 1995
;
Carabeo
et al
., 2003
). They also recruit the host’s rough endoplasmic
reticulum (rER), eventually resulting in a translocation of rER proteins into the inclusion
(
Dumoux
et al
., 2012
). Mitochondria are recruited to the inclusions of
C. psittaci
and
Waddlia chondrophila
(
Friis, 1972
;
Peterson and de la Maza, 1988
;
Matsumoto
et al
., 1991
;
Croxatto and Greub, 2010
).
Internalization, inclusion development and host-organelle recruitment are all mediated by
the secretion of effector proteins into the inclusion membrane and/or host cytoplasm by the
type III secretion (T3S) system (
Peters
et al
., 2007
). While the genes that encode this needle-
like secretion system are present in all chlamydial genomes (
Collingro
et al
., 2011
), T3S
structures have not been seen in environmental chlamydiae, and few structural details are
known about the T3S systems of pathogenic chlamydiae (
Matsumoto, 1979
;
Nichols
et al
.,
1985
;
Dumoux
et al
., 2012
).
Studying chlamydial cell biology is challenging because of their obligate intracellular
lifestyle and the lack of routine genetic tools (
Binet and Maurelli, 2009
;
Kari
et al
., 2011
;
Wang
et al
., 2011
;
Nguyen and Valdivia, 2012
). While many insights have come from
conventional electron microscopy (EM) studies, the chemical fixation, dehydration, plastic
embedding, thin sectioning and heavy-metal staining involved can lead to membrane
artefacts, misleading representations of the nucleoid structure or loss of entire cellular
components (
Pilhofer
et al
., 2010
). Here, we investigated three environmental chlamydiae,
Protochlamydia amoebophila
,
Parachlamydia acanthamoebae
and
Simkania negevensis
, by
electron cryotomography (ECT), which allows cells to be imaged in a near-native, ‘frozen-
hydrated’ state. This approach revealed not only new structural details of these obligate
intracellular bacteria at macromolecular resolution and in three dimensions but also provided
new perspectives on the bacteria-host interface.
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Results
Developmental stages of environmental chlamydiae
To investigate the ultrastructure of isolated cells, chlamydiae were purified from amoeba
cultures, plunge-frozen on EM grids and imaged intact. Twenty-five, twenty and twenty
tomograms were collected on purified
Simkania
,
Parachlamydia
and
Protochlamydia
cells
respectively. EBs and RBs could be distinguished by their size, morphology and the
granularity of their cytoplasm. EBs were coccoid and had diameters of 450 nm (
Simkania
,
±22,
n
= 9), 678 nm (
Parachlamydia
, ±32,
n
= 5) and 625 nm (
Protochlamydia
, ±170,
n
= 5)
(Fig. 1A, D and G). RBs were more pleomorphic and larger (667 nm,
Simkania
, ±46,
n
= 5;
838 nm,
Parachlamydia
, ±135,
n
= 4; 884 nm,
Protochlamydia
, ±88,
n
= 6) (Fig. 1C, F and
I). EBs exhibited regions of concentrated filamentous material (presumably condensed
DNA), with different texture from the rest of the EB cytoplasm. Smaller but otherwise
similar regions were also occasionally seen in RBs. Because of an irregularly shaped outer
membrane, the thickness of the RB periplasm was more variable than that seen in EBs.
Large numbers of ribosomes were found throughout the cytoplasm of both EBs and RBs
except in the region of the putatively condensed DNA within EBs. We also observed cells
with features characteristic of both developmental stages (Fig. 1B, E and H), including
multiple small patches of condensed DNA, probably representing intermediate stages in the
process of differentiation or redifferentiation.
Previous studies using conventional EM have reported that some chlamydiae exhibit
crescent shapes, and these ‘crescent bodies’ were suggested to represent an infectious life
stage of
Simkania
,
Parachlamydia
and
Protochlamydia
(
Greub and Raoult, 2002
;
Lamoth
and Greub, 2010
;
Nakamura
et al
., 2010
). Surprisingly, while our tomograms of intact as
well as cryosectioned cells (see later) allowed for the identification of EBs, RBs and
intermediate stages (Fig. 1), we never saw any crescent bodies (Fig. 1, Fig. 2A and B, Figs
3–5). We therefore explored if crescent bodies could be an artefact of chemical fixation and
dehydration/embedding.
Parachlamydia
cells were purified from asynchronous amoeba cultures and split into two
aliquots. One sample was processed with procedures similar to the original study describing
crescent bodies (
Greub and Raoult, 2002
): cells were fixed with glutaraldehyde and osmium
tetroxide, dehydrated, plastic-embedded, thin-sectioned, stained and imaged at room
temperature. Crescent bodies made up 47 ± 10% of all putative chlamydial cells (
n
= 813)
and had the typical shape and dimensions reported previously (
Greub and Raoult, 2002
)
(arrows in Fig. 2C and D). The second sample was treated in the same way, except that the
osmolarity and fixative concentration in the fixation buffer was reduced. Crescent bodies
were still present but only at a frequency of 2 ± 2% (
n
= 1355) (Fig. 2E). Osmolarity and
fixative concentration therefore influence the abundance of crescent bodies. This was further
supported by scanning EM (SEM) of purified chlamydial cells, where buffers with higher
osmolarity and higher fixative concentration also resulted in an increased percentage of
crescent bodies (Fig. 2F).
To distinguish between the effects of fixation and dehydration/plastic embedding, two
further aliquots of purified cells were cryopreserved (i.e. plunge-frozen) and imaged in a
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frozen-hydrated state in an electron cryomicroscope. No crescent bodies were found in
projection images of 584 cells, which were directly plunge-frozen after purification (Movie
S1). When cells were fixed with glutaraldehyde and osmium tetroxide before plunge-
freezing, crescent bodies were still absent (data not shown). Lysed cells were observed in
some cases, but none of those had a typical crescent shape or continuous intact membranes,
which are characteristic for crescent bodies. We conclude that crescent bodies are an artefact
of the combined effect of chemical fixation and dehydration/embedding.
Architecture of inclusions
Next chlamydiae were imaged inside their host. Because ECT is limited to thin (less than
~500 nm) samples, asynchronously infected amoeba cultures were pelleted, mixed with
cryoprotectant, high-pressure frozen, sectioned at cryotemperatures (150 nm section
thickness) and then imaged. Twenty-one, four and twenty-seven tomograms were collected
of vitreous cryosections of
Simkania
-,
Parachlamydia
- and
Protochlamydia
-infected
amoebae respectively. For comparison, we also collected tomograms of parallel samples
prepared by high-pressure freezing, freeze-substitution, plastic-embedding, thin-sectioning
and staining.
First, the localization of chlamydiae inside their host was investigated. Intracellular bacterial
cells were always seen surrounded by an inclusion membrane and never directly in the
cytoplasm (Figs 3 and 4, and Movies S2, S3 and S4). Single-celled inclusions and inclusions
packed with up to 17 chlamydial cells were observed in amoebae infected with
Simkania
(Fig. 3A–F) or
Parachlamydia
(Fig. 3G–L). RBs were the predominant stage within the
inclusions, some of them dividing by binary fission. In contrast, 92% (
n
= 52) of the
inclusions in amoebae infected with
Protochlamydia
contained a single bacterial cell (Fig.
4A, D and E). The 8% of inclusions harbouring more than one bacterium were not roundish
and tightly packed with bacteria, as seen for
Parachlamydia
and
Simkania
(Fig. 3); they
rather appeared like inclusions in the process of starting separation, with membranes
contracting in between the bacteria (Fig. 4B and C), or in a stage where two single-cell
inclusions were connected by a narrow membrane tube (the percentage of single-cell
inclusions in all cases might actually be slightly lower than noted, as extensions of the
inclusions above and below the section cannot be visualized).
Interestingly, we observed a difference in cell shape between
Simkania
EBs imaged inside
densely packed inclusions within their host versus after purification. While EBs imaged
inside these inclusions were frequently rod-shaped or elongated (cells labelled ‘EB’ in Figs
3F and 5A), purified EBs were always spherical (Fig. 1A).
Parachlamydia
and
Protochlamydia
EBs, in contrast, were always coccoid (Fig. 1D and G, and cells labelled
‘EB’ in Fig. 5B and C). RBs of all species had a somewhat polymorphic, spherical shape
inside the host cell.
Recruitment of ER by Simkania
Chlamydiae associate not only with the inclusion membrane, but some also recruit and
reshape entire host organelles. We found that inclusions of
Simkania
were always enveloped
by an additional membranous structure (Fig. 3A–F). The granularity inside the cisternae-like
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membrane sacs was different from the rest of the eukaryotic cytoplasm, suggesting that they
were part of a separate compartment. The endoplasmic reticulum (ER)-like membrane
architecture and the presence of many ribosomes on the cytoplasmic side of the distal
membrane identified the compartment as rER. Segmentations showed that the inclusions
were not entirely surrounded by the rER, however, leaving small patches of direct
connections between inclusion and amoeba cytoplasm (Fig. 3D). rER was found associated
with single-cellular (Fig. 3A, B and E) and multicellular inclusions filled with EBs and RBs
(Fig. 3C and F), indicating that ER recruitment occurs early after internalization and remains
throughout the intracellular stage.
In contrast with
Simkania
, no direct association of
Parachlamydia
or
Protochlamydia
inclusions and any host organelle was detected. Mitochondria were occasionally observed in
their vicinity (Figs 3G–L and 4A–E), but a specific co-localization was not suggested by
fluorescent labelling of mitochondria (Fig. S1).
Secretion systems
Translocation of chlamydial effector proteins into the inclusion membrane and into the host
cytoplasm is crucial for chlamydiae to shape their intracellular environment. In a
cryotomogram of a
Simkania
EB, we observed a structure with characteristics typical of a
T3S apparatus (Fig. 6A and B) (
Marlovits
et al
., 2004
). A density in the periplasm was
found to be similar to T3S basal bodies and connected to an extracellular needle-like
structure. The needle (length 63 nm, diameter 9 nm) seemed to be engaged with a
membranous structure, possibly a remnant of the host cell. The dimensions of the apparatus
were similar to projections seen on the surface of infectious
C. psittaci
cells (
Matsumoto,
1979
). Interestingly, the otherwise relatively narrow distance between the inner and outer
membrane (13 nm) in
Simkania
EBs required a bulging (41 nm) of the cytoplasmic
membrane to accommodate the basal body. Such widening of the periplasm was observed
frequently in the same and other EBs (Figs 6A and S2). Basal body-like densities inside
these bulges indicate that they likely represented T3S structures as well, but the
corresponding needles were probably sheared off during purification. A pronounced
widening of the periplasmic space was also reported for T3S structures of
C. trachomatis
(
Dumoux
et al
., 2012
).
Putative T3S systems in
Parachlamydia
showed a similar bulging of the periplasm in the
region of the basal body (36–44 nm rather than 17 nm) (Fig. 6C–F). The needle structure
was substantially different compared with
Simkania
, with a length of 38–42 nm, a diameter
of 6–7 nm and a widening (12 nm) of the needle 7 nm from its tip (Fig. 6D and F). A T3S-
like structure found on a
Protochlamydia
EB comprised a 52-nm-long, 7-nm-diameter
needle, and may have also had a widening close to the needle tip (Fig. 6G and H).
Periplasmic bulging was not observed, however, as the width of the periplasm in
Protochlamydia
was already ~40 nm.
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Discussion
EBs and RBs in a life-like state
Early conventional EM studies suggested that the DNA in EBs is condensed (
Moulder,
1966
), which was later found to be mediated by histone-like proteins (
Barry
et al
., 1992
).
However, nucleoid structure in particular is prone to artefacts introduced by fixation,
dehydration and staining in conventional EM (
Pilhofer
et al
., 2010
). While a more recent
ECT study imaged
C. trachomatis
cells preserved in a near-native, frozen-hydrated state,
such ultrastructural details were unfortunately not resolved probably due to instrumental
limitations (
Huang
et al
., 2010
). Again plunge-freezing cells, but using higher electron
energies and energy filtration, here we have confirmed that EB genomes are indeed densely
packed. Our finding of ribosomes in EBs is consistent with the notion that they are
metabolically active to some degree (
Haider
et al
., 2010
;
Sixt
et al
., 2013
) rather than
completely dormant (
Hatch
et al
., 1985
). Besides size differences of EBs and RBs not
observed before for environmental chlamydiae, another distinguishing feature between the
developmental stages we noted was the variable periplasmic width in RBs. This is consistent
with the notion that lower abundances of stabilizing cysteine-rich proteins in RBs result in
more flexible outer membranes (
Hatch
et al
., 1986
). Similarly, the more flexible shape and
deformation of
Simkania
EBs inside host cells compared with
Protochlamydia
and
Parachlamydia
might be a consequence of differences in cell envelope architecture, such as
the absence of the cysteine-rich proteins that
Simkania
lacks in contrast with all other
chlamydiae (
Collingro
et al
., 2011
).
Crescent bodies are an artefact
While crescent-shaped cells had been seen previously and thought to represent a distinct
developmental stage (
Greub and Raoult, 2002
;
Lamoth and Greub, 2010
;
Nakamura
et al
.,
2010
), here we showed that they are artefacts of conventional EM methods. While no
crescent bodies have been reported for the pathogenic
Chlamydiaceae
, EBs with peculiar
stellate outlines were found occasionally. The previous hypothesis that this morphology
could be attributed to EM preparation methods as well (
Matsumoto, 1988
) is supported by
our results. The reason for crescent bodies not being observed in
Chlamydiaceae
could be
differences in their outer membrane protein composition compared with environmental
chlamydiae (
Heinz
et al
., 2009
;
Collingro
et al
., 2011
), leading to different effects during
chemical fixation and dehydration/embedding.
Trapezoidal, dumbbell-shaped and elongated intracellular
Simkania
EBs have also been
described in the past (
Kahane
et al
., 2001
;
Michel
et al
., 2005
;
Henning
et al
., 2007
). While
we found elongated morphologies especially in cells tightly packed in inclusions,
trapezoidal and dumbbell-shaped forms were never seen, suggesting that those are also
artefacts. Conventional EM studies of other environmental chlamydiae EBs have reported
head-and-tail, star and rod shapes (
Kostanjsek
et al
., 2004
;
Karlsen
et al
., 2008
;
Lienard
et
al
., 2011
). It remains unclear whether these morphologies are natural.
Shapes of bacteria from other phyla have also been reported to be affected by chemical
fixation and dehydration. For instance, crescent-shaped cells were observed for
Gemmata
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obscuriglobus
(
Lindsay
et al
., 1995
), a member of the chlamydial sister-phylum
Planctomycetes
, and the mollicute
Acholeplasma ladlawii
(
Lemcke, 1972
), showing that
fixation conditions must be chosen carefully to preserve the cell shape and that the
description of new shapes based on fixed cells should be handled with caution.
Diversity of the intracellular niche of environmental chlamydiae
Most chlamydiae are known to reside inside the host-derived membranous inclusion after
host cell invasion (
Hackstadt
et al
., 1997
), but
Parachlamydia
and
Simkania
have also been
reported to be localized directly in the cytoplasm (
Michel
et al
., 1994
;
Greub and Raoult,
2002
). Here, intracellular chlamydiae were always seen surrounded by an inclusion
membrane, supporting the importance of this host-bacterium interface for intracellular
survival and replication. Inclusions in
Protochlamydia
infections were exclusively
unicellular, but
Simkania
and
Parachlamydia
cells were more commonly found in
multicellular inclusions.
Some chlamydial species are known to recruit and reshape entire host organelles including
mitochondria, Golgi stacks or the ER (
Peterson and de la Maza, 1988
;
Matsumoto
et al
.,
1991
;
Heuer
et al
., 2009
;
Croxatto and Greub, 2010
;
Dumoux
et al
., 2012
). We found that
Simkania
inclusions are almost entirely enveloped by the rER (Fig. 3), adding additional
layers to the host-bacterial interface. In this way,
Simkania
might use a similar strategy as
the facultative intracellular pathogens
Legionella pneumophila
and
Brucella abortus
(
Swanson and Isberg, 1995
;
Abu Kwaik
et al
., 1998
;
Roy, 2002
). However, in contrast with
L. pneumophila
and
B. abortus
phagosomes, the
Simkania
inclusion does not fuse with ER-
derived vesicles, and
Simkania
thus remains inside the inclusion. The tight association of the
Simkania
inclusion with the ER could nevertheless provide similar benefits such as
prevention from fusing with lysosomes. Interestingly, the abilities to recruit ER and to
replicate in human and insect cells coincide in
Simkania
(
Kahane
et al
., 2007
;
Sixt
et al
.,
2012
) and members of the pathogenic chlamydiae (
Dumoux
et al
., 2012
), but are absent in
Parachlamydia
and
Protochlamydia
.
Species-specific differences in inclusion morphology and recruitment of host organelles are
likely due to the presence of different effector proteins in the inclusion membrane (
Rockey
et al
., 1997
;
Betts
et al
., 2009
;
Heinz
et al
., 2010
). Adaptation to different hosts likely drove
the diversification of environmental chlamydiae (
Bertelli
et al
., 2010
;
Collingro
et al
., 2011
).
T3S systems
Translocation of chlamydial effector proteins through the elaborate cell envelope and the
inclusion membrane requires a secretion system and is thought to be accomplished by the
T3S system. T3S systems are encoded in all known chlamydial genomes (
Peters
et al
., 2007
;
Collingro
et al
., 2011
), and T3S proteins were detected during all stages of infection in
members of the pathogenic chlamydiae (
Fields
et al
., 2003
). For the first time, we detected
T3S-like structures in environmental chlamydiae, providing evidence for its conservation
and crucial role in the infectious life cycle of modern and likely ancient chlamydiae. Fewer
T3S-like structures were observed by ECT in environmental chlamydiae than by
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conventional transmission electron microscopy in pathogenic chlamydiae (
Matsumoto,
1982
;
Wilson
et al
., 2006
).
T3S needle tip proteins in other bacteria are known to be highly adapted to the host (
Abby
and Rocha, 2012
). It has been unclear whether the chlamydial T3S needle harbours a tip
protein at all. To date, only one candidate for a chlamydial needle tip protein has been
identified, but it remains unclear whether it rather functions as an effector (
Markham
et al
.,
2009
;
Stone
et al
., 2012
). The subterminal widening of the needle in
Parachlamydia
and
Protochlamydia
seen here indicates that the chlamydial T3S apparatus likely does include a
needle tip protein. Interestingly, T3S structures were not seen on purified or cryosectioned
RBs perhaps because the juxtaposition of the RB outer membrane and inclusion membrane
effects the length of the needle.
Imaging bacteria–host interactions in a near-native state
Finally, this is the first study to image bacteria inside their host in a near-native, frozen-
hydrated state. In addition to avoiding and uncovering artefacts, this approach provided
novel insights into the nature of the host-bacterial interface. Because amoebae can also serve
as hosts for important pathogens such as
Legionella pneumophila
,
Vibrio cholerae
,
mycobacteria,
Francisella tularensis
,
Pseudomonas aeruginosa
and
Helicobacter pylori
as
well as bacterial symbionts like
Amoebophilus asiaticus
,
Paracaedibacter symbiosus
or
Procabacter acanthamoebae
(
Horn and Wagner, 2004
;
Schmitz-Esser
et al
., 2008
), our
approach should prove helpful in the study of many other important bacteria–host
interactions in the future.
Experimental procedures
Cultivation of organisms and staining of mitochondria
Acanthamoeba castellanii
UWC1 infected with
Parachlamydia acanthamoebae
UV7 or
Simkania negevensis
and
A. castellanii
Neff infected with
Protochlamydia amoebophila
UWE25 were cultivated in TSY (trypticase soy broth with yeast extract) medium (30 g l
−1
trypticase soy broth, 10 g l
−1
yeast extract, pH 7.3) at 20°C. Amoebal growth was monitored
by light microscopy and medium was exchanged every 3–6 days. The presence and identity
of the chlamydial symbionts was checked regularly by fluorescence
in situ
hybridization
(FISH) combined with 4
′
,6-diamidino-2-phenylindole staining of infected cultures using
specific probes for the respective symbiont as described previously (
Schmitz-Esser
et al
.,
2008
). In addition, the identity of the symbionts was verified by isolation of DNA from
cultures followed by amplification and sequencing of the 16S rRNA genes. For staining of
mitochondria,
A. castellanii
infected with chlamydial symbionts were incubated with 2 μM
MitoTracker Orange CMTMRos (Molecular Probes) in TSY for 45 min. Cells were fixed
with 4% paraformaldehyde, followed by FISH with specific probes.
Purification of chlamydiae
Infected
A. castellanii
cultures were harvested by centrifugation (7197 ×
g
, 10 min), washed
in Page’s amoebic saline (PAS) (
Page, 1976
), centrifuged and resuspended in PAS. Amoeba
cells were ruptured by vortexing with an equal volume of glass beads for 3 min. Glass beads
Pilhofer et al.
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and cell debris were removed by centrifugation (5 min, 300 ×
g
). The supernatant was
filtered through a 1.2 μm filter and centrifuged at maximum speed for 10 min. The obtained
pellet was resuspended in PAS.
Conventional transmission EM
To analyse the impact of fixation and to compare the effect of different fixation buffers on
the morphology of
Parachlamydia
, chlamydiae were purified from their amoeba hosts, and
the sample was divided into three parts. One part was immediately plunge-frozen (see later).
The second part was fixed in 4% glutaraldehyde in phosphate buffered saline (PBS, 130 mM
NaCl, 10 mM Na
x
PO
4
; pH 7.2–7.4) for 1 h, washed in PBS and further fixed in 1% osmium
tetroxide in PBS for 1 h followed by two washing steps. The third part was fixed in the same
way as the second sample, except that 2% glutaraldehyde in phosphate buffer (10 mM
Na
x
PO
4
; pH 7.2–7.4) was used as first fixative and that the 10 mM phosphate buffer
replaced PBS in the following washing and fixation steps. Samples were dehydrated in
ethanol and acetone through a graded series, embedded in Epon-Araldite (Electron
Microscopy Sciences, Port Washington, PA), thin-sectioned with a UC6 ultramicrotome
(Leica, Vienna, Austria), and stained with uranyl acetate and lead citrate. Two-dimensional
images were recorded on a Tecnai T12 TEM (FEI, Eindhoven, the Netherlands).
For room temperature EM of high-pressure frozen/freeze substituted samples, infected
amoeba cells were high-pressure frozen (see later). The frozen domes were transferred under
liquid nitrogen to cryotubes containing 2% or 0.04% glutaraldehyde in acetone. The tubes
were placed in a model AFS freeze-substitution machine (Leica) and freeze-substituted at
−90°C for 60 h, then warmed to −20°C over 10 h. Cells were rinsed 3× with cold acetone,
then post-fixed with 2.5% osmium tetroxide in acetone at −20°C for 24 h. The samples were
then warmed to 4°C over 2 h, rinsed 3× with cold acetone, and embedded in Epon-Araldite
resin (Electron Microscopy Sciences). Following polymerization, semi-thin (200 nm)
sections were cut with a UC6 ultramicrotome (Leica) and placed on Formvar-coated, copper/
rhodium 1 mm slot grids (Electron Microscopy Sciences). Sections were stained with uranyl
acetate and lead citrate, and imaged in a Tecnai T12 TEM (FEI). Dual-axis tilt-series were
acquired using SerialEM (
Mastronarde, 2005
), then subsequently calculated and analysed
using IMOD (
Kremer
et al
., 1996
) on an Apple MacPro computer.
Plunge-freezing
For plunge-freezing, copper/rhodium EM grids (R2/2 or R2/1, Quantifoil, Jena, Germany)
were glow-discharged for 1 min. A 20×-concentrated bovine serum albumin-treated solution
of 10 nm colloidal gold (Sigma, St Louis, MO) was added to purified chlamydiae (1:4 v/v)
immediately before plunge freezing. A 4 μl droplet of the mixture was applied to the EM
grid, then automatically blotted and plunge-frozen into a liquid ethane-propane mixture
(
Tivol
et al
., 2008
) using a Vitrobot (FEI) (
Iancu
et al
., 2006
).
Cryosectioning
Acanthamoeba castellanii
cells continuously infected with either
Simkania
,
Parachlamydia
or
Protochlamydia
were mixed with uninfected amoeba cells at a ratio of 1:1 and incubated
for 24 h. For
Parachlamydia
, the ratio of infected to uninfected cells was 5:1. Amoebae were
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harvested (7197 ×
g
, 10 min), and the pellet was mixed with 40% dextran (w/v) in PAS. The
samples were transferred to brass planchettes and rapidly frozen in a HPM010 high-pressure
freezing machine (Bal-Tec, Leica). Cryosectioning of the vitrified samples was done as
previously described (
Ladinsky
et al
., 2006
;
Ladinsky, 2010
). Semi-thin (90–200 nm)
cryosections were cut at −145°C or −160°C with a 25°Cryo diamond knife (Diatome, Biel,
Switzerland), transferred to grids (continuous carbon-coated 200-mesh copper grids or 700-
mesh uncoated copper grids) and stored in liquid nitrogen.
ECT
Images were collected using a Polara 300 kV FEG transmission electron microscope (FEI)
equipped with an energy filter (slit width 20 eV; Gatan, Pleasanton, CA) on a lens-coupled 4
k × 4 k UltraCam charge-coupled device (CCD) (Gatan) or K2 Summit direct electron
detector (Gatan). Pixels on the CCD represented 0.95 nm (22 500×) or 0.63 nm (34 000×) at
the specimen level. Typically, tilt series were recorded from −60° to +60° with an increment
of 1° at 10 μm under-focus. The cumulative dose of a tilt-series was 180–220 e−/Å2 (for
whole cells) or 100–150 e−/Å2 (for cryosections). UCSFTOMO (
Zheng
et al
., 2007
) was
used for automatic acquisition of tilt-series and two-dimensional projection images. Three-
dimensional reconstructions were calculated using the IMOD software package (
Kremer
et
al
., 1996
) or Raptor (
Amat
et al
., 2008
). Tomograms of cryosections were reconstructed
using IMOD’s patch tracking to generate the aligned stack (
Kremer
et al
., 1996
).
Tomograms were visualized and segmented using 3dMOD (
Kremer
et al
., 1996
).
SEM
Glass coverslips (12-mm diameter) were cleaned in acidic ethanol, dried for 1 h at 60°C and
coated with 0.01% poly-L-Lysine solution for 10 min. Two hundred microlitres of purified
Parachlamydia
in the respective buffer were spotted onto the dry coverslip. After 10 min
non-attached cells were removed, and remaining cells were fixed for 1 h at room
temperature using the following fixatives: 2% glutaraldehyde in 10 mM phosphate buffer
(pH 7.2), 2.5% glutaraldehyde in 3 mM cacodylate buffer (pH 7.2), 2% glutaraldehyde in
DGM-21A-defined medium (
Haider
et al
., 2010
), 4% glutaraldehyde in 10 mM phosphate
buffer with 130 mM NaCl, and 4% glutaraldehyde in 10 mM phosphate buffer with 260 mM
NaCl. After three washing steps (5 min each) in the respective buffer, cells were further
fixed in 1% osmium tetroxide in the respective buffer for 1 h at room temperature and
washed again three times. Samples were dehydrated in acetone and chemically dried in
hexamethyldisilazane. Glass slides were gold coated for 160 s using default settings (Agar
sputter coater B7340) and analysed using a Philips XL-30 ESEM. For analysis, 10 or more
random SEM images with 36 or more individual putative bacterial cells in total were taken.
Roundish or crescent-shaped objects with a diameter of 0.5–1 μm were counted as bacterial
cells. Each cell was then classified into one out of four morphological types (crescent shape,
large invaginations, small invaginations, coccoid), and the percentage of each type was
determined. Osmolarity measurements of buffers and fixatives were performed using an
Advanced Micro 3MO plus osmometer (Block Scientific, New York, NY). Samples and
standards were measured three times each.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
Acknowledgments
This work was funded by the Austrian Science Fund FWF (Y277-B03 to MH), the European Research Council
(ERC StG ‘EvoChlamy’ to MH), the Caltech Center for Environmental Microbiology Interactions (to GJJ, MP),
and a gift from the Gordon and Betty Moore Foundation to Caltech.
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