Chlamydia, escaping a fate worse than death

Markus C. Kerr1, Guillermo A. Gomez1, Charles Ferguson1, Maria C. Tanzer2,3, James

M. Murphy2,3, Alpha S. Yap1, Robert G. Parton1, Wilhelmina M. Huston 4 and Rohan

D. Teasdale1*

AFFILIATIONS:

1. Institute for Molecular Bioscience, The University of Queensland, St Lucia,

Queensland, Australia.

2. he Walter and Eliza Hall Institute of Medical Research, Parkville, Victoria,

Australia.

Department of Medical Biology, University of Melbourne, Parkville, Victoria,

Australia.

4. i3 Institute, University of Technology, Sydney, New South Wales, Australia.

(*) CORRESPONDING AUTHOR:

Rohan Teasdale, PhD

Institute for Molecular Bioscience

The University of Queensland

St. Lucia, QLD 4072

AUSTRALIA

E-MAIL: R.Teasdale@uq.edu.au

PHONE: 61-7-3346-2056

RUNNING TITLE: Calpains mediate chlamydial egress

HIGHLIGHTS:

Chlamydial egress is primarily mediated by the host.

Chlamydia-induced cell death is independent of BAK, BAX, RIP1 and

caspase activity.

Calpains regulate chlamydial inclusion rupture required for chlamydia

egression but not host cell death.

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Summary

Remarkably little is known about how intracellular pathogens exit the host cell in

order to infect new hosts. Pathogenic Chlamydia spp. egress by first rupturing their

replicative niche, the inclusion, before rapidly lysing the host cell. We developed a

unique laser ablation strategy to specifically disrupt the chlamydial inclusion thereby

uncoupling inclusion rupture from the subsequent cell lysis. Using this approach we

define a novel role for the host cell calpains in inclusion rupture, but not subsequent

cell lysis. Further, with genome-editing technologies we demonstrate that chlamydial

egress is mediated by a rapid non-apoptotic non-necroptotic cell death pathway

independent of BAK-, BAX-, RIP1- and caspase-activity. Both inclusion rupture and

rapid host cell death work in concert to efficiently liberate the pathogen from the host

cytoplasm, thereby minimising exposure to damaging reactive oxygen species,

promoting subsequent secondary infection and in doing so reconcile the pathogen's

established capacity to promote cell survival and induce cell death as required.

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Introduction

Chlamydia trachomatis is the most prevalent bacterial sexually transmitted infection

amongst humans and is the leading cause of infectious blindness worldwide. As an

obligate intracellular pathogen, Chlamydia maintains exquisite control over an

assortment of host cellular processes during its dimorphic infectious cycle. Most

prominent amongst these is the formation of an intracellular replicative niche from the

host cell’s membrane trafficking pathways (Elwell and Engel, 2012) and the profound

pro-survival influence its presence elicits during the replicative phase of infection

(Fan et al., 1998; Sharma and Rudel, 2009).

Chlamydia invades host cells as a non-replicative Elementary Body (EB) through the

action of a Type 3 Secretion System (T3SS) that serves to deliver bacterial effector

molecules to modulate the host’s membrane trafficking and cytoskeletal elements.

Once intracellular, Chlamydia alters the encompassing vacuole to create its replicative

niche, called an inclusion, where it transitions into its metabolically active replicative

Reticulate Body (RB) form. During the later stages of the pathogen’s life-cycle,

Chlamydia asynchronously transforms back into its EB form before it egresses from

the cell in one of two mechanisms. Pioneering work by Hybiske and Stephens (2007)

reported that the chlamydial inclusion is either extruded intact from the cell or

ruptures immediately prior to cell lysis (Hybiske and Stephens, 2007). The extrusion

mechanism is an actin-dependent process (Hybiske and Stephens, 2007) recently

reported to be coordinated by the actions of myosin phosphatase, myosin light chain 2

(MLC2), myosin light chain kinase (MLCK), and myosin IIA and IIB (Lutter et al.,

2013) and septins (Volceanov et al., 2014).

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Whilst extrusion is speculated to contribute to evasion of the host immune response

and long-distance dissemination, release of the EBs to infect new host cells ultimately

necessitates lysis of both the inclusion and limiting membrane of the cell and/or

extrusion. Hybiske and Stephens (2007) demonstrated the requirement for cysteine

protease activity during rupture of the inclusion using the pan-cysteine protease

inhibitor E-64 and intracellular calcium for the subsequent lysis of the limiting

membrane and release of the Chlamydia into the extracellular milieu (Hybiske and

Stephens, 2007). The asynchronous nature of chlamydial egress has, however,

impeded further dissection of the process and remarkably little is known about the

molecular events involved, particularly the identity of the cysteine proteases involved

in inclusion rupture. Here we have applied a novel laser-ablation approach to dissect

the molecular events regulating release of the pathogen following inclusion rupture.

Whilst maintaining its established anti-apoptotic influence upon the cell, chlamydial

inclusion rupture triggers a programmed necrotic cell death pathway, that rapidly

liberates the pathogen from the destructive cytoplasm. This process is triggered

independent of the stage of infection and in the presence of inhibitors of bacterial

protein synthesis suggesting that it is regulated by the host rather than orchestrated by

the pathogen. We reveal a novel role for host calpains in inclusion rupture but not cell

lysis highlighting the host’s pivotal role in regulating this process.

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Results

Inclusion rupture is directly coupled to chlamydial egress

Hybiske and Stephens (2007) demonstrated that the chlamydial inclusion ruptures

immediately prior to cell lysis during the later stages of infection (Hybiske and

Stephens, 2007). We recapitulate this observation in a HeLa reporter cell line stably

expressing mCherry-tagged Rab25 to monitor the integrity of Chlamydia trachomatis

strain LGVII (CTL2) inclusions throughout the infection (Teo et al., 2016), CFP-

Histone 2-B to highlight the nuclei and soluble GFP to mark the cytoplasm. From ~36

hours post-infection (h p.i.) the inclusions of infected cells begin to rupture, denoted

by the loss of inclusion integrity and influx of cytoplasmic GFP into the inclusion

lumen (asterisk) leading to an overall dimming of the GFP-fluorescence, in an

asynchronous manner before loss of cellular plasma membrane integrity from 15-

30min post-inclusion rupture (Fig. 1A). Notably the nuclei of cells following

inclusion rupture general maintain their overall structure, condensing moderately prior

to cell lysis (arrows). Although the kinetics of the lytic process post-rupture were

extremely consistent, inclusion rupture was observed in a stochastic manner anywhere

from 36-72h p.i. Whilst it is recognised that cysteine protease activity is required for

inclusion rupture and intracellular calcium signalling is intrinsic to subsequent cell

lysis (Hybiske and Stephens, 2007) the manifest asynchronous nature of inclusion

rupture has proven refractory to more detailed investigation of the molecular events

involved in chlamydial egress. Here we have applied a unique 2-photon ablation

system (Gomez et al., 2015) to specifically rupture the chlamydial inclusion (reticle)

without damaging the plasma membrane of the cell thereby allowing us to reliably

interrogate the process in high resolution and uncouple inclusion rupture from

subsequent cell lysis (Fig. 1B).

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HeLa cells stably expressing mCherry-Rab25 were seeded onto imaging plates,

transfected with soluble GFP and infected with CTL2 for 48h p.i. The cells were

imaged live on a line scanning confocal microscope for 3 frames before a 0.6μm3

region of the inclusion was ablated (reticule) and the samples imaged further (Fig.

1B). Consistent with native egress, immediately following ablation the inclusion is

observed to fill with soluble GFP before it is observed to collapse in a manner similar

to that observed previously using mechanical disruption (Heinzen and Hackstadt,

1997). Maintenance of the plasma membrane’s integrity during the ablation was

affirmed as the soluble GFP was observed to fill the inclusion (leading to an overall

dimming) but remain within the cell. From 15-30 minutes following inclusion rupture,

ablated cells are observed to bleb extensively, before contracting and releasing the

contents of their cytoplasm into the extracellular milieu. Notably, throughout this

process the overall structure of the nucleus is maintained with some degree of

condensation similar to that observed during native lysis observed (Fig. 2B, arrows).

Surprisingly, ablation of inclusions in HeLa cells infected for 24h p.i. with GFP-

expressing CTL2 also triggered cell death and lysis (Fig. 1C). Whilst the initiation of

cell death was modestly delayed, once initiated, the cells appeared to die with similar

kinetics to those ablated at 48h p.i. suggesting that the cell death pathway involved is

not dependent upon stage of infection. The consequences of inclusion ablation upon

cell survival were quantified for 10 cells in the presence of cellular and bacterial

inhibitors and at different stages of infection (Fig 1D). As in native egress, post-

inclusion ablation-triggered lysis of the cell was sensitive to intracellular calcium

signalling (Hybiske and Stephens, 2007) as ablation of inclusions in 24h p.i. cells pre-

cultured for 1h in calcium-free Ringer's solution supplemented with 1,2-bis(2-

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aminophenoxy)ethane-N,N,N′,N′-tetraacetate (BAPTA)-acetoxymethyl (-AM) ester

resulted in sustained viability of cells (Fig 1D & E). Bacterial protein synthesis was

not required to trigger cell death as treatment of cells with 100μg/ml chloramphenicol

from 24h p.i. did not impact upon ablation-triggered cell death response, nor did the

stage of infection, suggesting that it is the host that drives the lytic process rather than

the pathogen (Fig. 1D). Notably it was recently suggested that the secreted chlamydial

protease-like activity factor (CPAF) accumulates within the chlamydial inclusion and

is released upon rupture so that it may target vimentin intermediary filaments and

components of the nuclear envelope for degradation (Snavely et al., 2014). Live

imaging of 2xGFP-tagged vimentin following laser-mediated inclusion rupture

recapitulated this observation with a rapid dissolution of filamentous structures and a

rapid translocation of 2xGFP-tagged vimentin into the nucleus. 2xGFP-vimentin

dissolution was blocked in the presence of a CPAF inhibitor (PMID: 21767809)

following inclusion rupture (Supplementary Fig. 1). Consistent with observations

made using a CPAF-null chlamydial strain (Snavely et al., 2014) inhibition of CPAF

activity did not block cell lysis following inclusion rupture.

Thus the lytic cell death observed subsequent to inclusion ablation mirrors the kinetics

and has the molecular features observed during native inclusion rupture. Surprisingly,

however, cell lysis occurs regardless of the stage of infection at which the inclusion is

disrupted and in the presence of bacterial protein synthesis and protease inhibitors

suggesting that the mechanism by which the cell death occurs and therefore

subsequent chlamydial egress is primarily coordinated by the host (Fig 1D).

Importantly, the specificity of the induced cell death response to rupture of the

inclusion is evinced by the observation that ablation of mCherry-Rab25-labelled

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endosomal membranes swollen in the presence of the endosomal trafficking inhibitor

YM201636 (Jefferies et al., 2008), in both infected and uninfected cells, does not

trigger the cell death response (Fig. 1F). Laser-ablation of chlamydial inclusions

therefore represents a directed means to trigger chlamydial egress in a controlled

fashion uncoupled from the contribution of the pathogen itself. Furthermore, inclusion

ablation mid-way through the intracellular development of the pathogen allows one to

examine the subsequent molecular events involved in high resolution without the

confounding impact of the profound metabolic burden or morphological distortion the

pathogen places upon the cell at the later stages of infection.

Premature inclusion rupture is detrimental to chlamydial development

Our system provides unparalleled spatial and temporal resolution and the means to

examine the consequences of inclusion rupture at the individual infected cell and

subcellular level in isolation. Initially correlative light and electron microscopy was

employed to compare the intraluminal environment of intact and ablated inclusions

between and within infected cells thereby providing internal controls for the protocol.

Once again, time-lapse videomicroscopy revealed the rapid condensation and

immobilisation of ablated 24h p.i. inclusions when compared with intact inclusions

prior to fixation 10 minutes post-ablation and processing for transmission electron

microscopy (Fig. 2A). Electron micrographs of these particular inclusions

demonstrated that, whilst intact inclusions present characteristic spacious arrangement

of chlamydial RBs in an electron-lucent lumen (Fig. 2B cell 1), the RBs of ablated

inclusions are compacted against one another in a tight arrangement with cytosol

distributed between them (Fig. 2B cell 2). Intriguingly, there was also evidence of RB

swelling in the ablated inclusions suggestive of a destructive impact of inclusion

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rupture upon the pathogen (Fig. 2B cell 2, arrows). Given that the lumen of the

chlamydial inclusion shares most biophysical properties in common with the

cytoplasm and there is a free exchange of cytoplasmic ions (Grieshaber et al., 2002),

we considered whether this bactericidal impact may reflect exposure to typically

excluded reaction oxygen species (ROSs) now freely accessing the bacteria from the

cytoplasm. Indeed, infection with C. trachomatis is recognised to induce the

production of ROSs early in infection with subsequent inaction of the host’s NADPH

oxidase suggesting that the pathogen actively suppresses ROS production to promote

infection (Boncompain et al., 2010). Paradoxically, inhibiting ROS production during

chlamydial infections also supresses the growth of the bacteria suggesting that the

pathogen requires host-derived ROS for optimal growth (Abdul-Sater et al., 2010). To

examine this more directly, CTL2-infected mCherry-Rab25 cells were imaged live

and the inclusion membranes ablated 24 h p.i. in the presence of CellROX® Green, a

fluorogenic probe that exhibits bright fluorescence upon oxidation by ROSs. It was

immediately apparent that the ROS probe did not detect and ROSs within the

chlamydial inclusion (Fig. 2C & 2D). Strikingly, ablation of the inclusion yielded a

dramatic accumulation of CellROX® Green fluorescence within the inclusion

indicating oxidation by ROSs. This was confirmed using the antioxidant, N-

acetylcysteine to quench ROS-mediated oxidation in ablated cells (data not shown).

ROSs are generated as by-products of cellular metabolism, primarily in the

mitochondria (Starkov, 2008). We used our ablation system to monitor the

mitochondria of cells immediately following inclusion rupture (Fig. 2E). Strikingly,

we observed swelling and eventual dimming of Mitotracker® Red CMXRos-stained

mitochondria (arrows) following inclusion ablation suggesting a transition in

mitochondrial permeability. In addition, we observed accumulation of the

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Mitotracker® fluorescence within the inclusion following inclusion rupture indicating

the now ready access of the thiol-reactive dye to the pathogen once the inclusion

membrane is compromised. Thus we now provide direct evidence that the chlamydial

inclusion provides a protective barrier to bactericidal concentrations of cytosolic

ROSs and that inclusion rupture triggers mitochondrial swelling, likely elevating

cytosolic ROSs concentrations still further. Exposure to ROSs clearly damages the

delicate RBs, and possibly also the more robust EBs, hence necessitating the rapid

release of the pathogen through a lytic cell death pathway to ensure viability of the

infectious progeny.

Inclusion rupture-induced Cell Death is not apoptotic

The calcium-dependent and highly coordinated nature of the rupture-induced cell lysis

is suggestive of a Programmed Cell Death (PCD) pathway (Fuchs and Steller, 2015;

Scorrano et al., 2003). There are, however, discordant views on the contribution of

PCDs in chlamydial infection biology. Whilst Vats and colleagues suggest that

caspase-dependent PCD, or classical apoptosis, is prominent amongst Chlamydia-

infected primary cervical epithelial cells (Vats et al., 2010), Schoier et al (2001) report

that apoptosis does not appear to be the primary mode of death for infected cells

within in vitro culture systems, rather it is prominent amongst the neighbouring

uninfected cells (Schoier et al., 2001). We demonstrate that the nuclear ultrastructure

of cells in which inclusions have ruptured appear to be, for the most part,

indistinguishable from those with intact inclusions (compare nuclei of cell 1 with cells

2 & 3 of Fig. 2B) lacking the characteristic condensation and peripheralisation of

chromatin (pyknosis) observed in classically apoptotic cells. They do, however,

present blebbing of the plasma membrane, cleavage of vimentin (Supplementary Fig.

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1) and swelling of mitochondria (Fig. 2E), which are all prominent features of

apoptosis (Byun et al., 2001). This prompted us to investigate the mechanism by

which the cells were dying more thoroughly.

The caspases are a family of cysteine proteases integral to apoptosis and cysteine

protease activity has been demonstrated to be required for inclusion rupture and

consequently chlamydial egress (Hybiske and Stephens, 2007). Our laser-mediated

approach also allows us to uncouple the events both prior to and following inclusion

rupture with fidelity. Among the caspases, caspase 3 is a frequently activated death

protease catalysing the cleavage of many cellular proteins. To monitor caspase 3

activity in cells in which the inclusion had been ruptured we initially employed a

Förster Resonance Energy Transfer (FRET)-based reporter sensor that encodes a

caspase 3 cleavage linker sequence DEVD, Casper3-GR (Evrogen). Following

ablation of the chlamydial inclusion, cells presented sustained FRET signal

suggesting that that the caspase 3 cleavage linker was maintained (Fig. 3A). Under the

same imaging conditions, a striking loss in FRET signal was observed following

induction of apoptosis with 100ng/ml TNFα and 10μg/ml cycloheximide for 4h (data

not shown) demonstrating the efficacy of the commercial probe. Furthermore,

treatment of cells with the pan-caspase inhibitor carbobenzoxy-valyl-alanyl-aspartyl-

[O-methyl]- fluoromethylketone (Z-Vad-fmk, 50μM) did not inhibit ablation-

triggered cell death (Fig 3B) or native egress of the pathogen (Fig. 3C). Similarly,

treatment with the caspase 1 selective inhibitor, VX-765, did not inhibit inclusion

rupture-triggered death (Supplementary Fig. 2A) or native egress of the pathogen

(Supplementary Fig. 2B).

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The Bcl-2 family of proteins constitute critical control points in the intrinsic and

extrinsic apoptotic pathways (Danial and Korsmeyer, 2004). Ordinarily, pro-apoptotic

BAX resides in the cytosol or is loosely associated with membranes but, in response

to death signals, is inserted into the mitochondrial outer membrane as a homo-

oligomerised multimer resulting in mitochondrial dysfunction. Similarly, in healthy

cells, an interaction between the voltage-dependent anion channel protein 2 (VDAC2)

with inactive pro-apoptotic BAK keeps this lethal molecule in check at the

mitochondrion. In response to stress signals, this interaction is displaced allowing

BAK to homo-oligomerise causing mitchondrial dysfunction. In both cases, formation

of the so-called Mitochondrial Outer Membrane Pore (MOMP) results in leakage of

mitochondrial content which in turn activates initiator and downstream effector

caspases. To definitively rule out apoptosis and to examine whether the observed

mitochondrial swelling was a consequence of BAK and/or BAX activity, we applied

CRISPr technology to block formation of the MOMP by selectively disrupting BAK

and/or BAX in HeLa cells (Figure 3D). Cells knocked out for BAK and BAX are

established to be intrinsically resistant to be intrinsic and extrinsic apoptotic stimuli as

well as selected necrotic stimuli (Karch et al., 2013). Consistent with these

observations our BAK/BAX double-knockout cells as well as those treated with Z-

Vad-fmk showed complete insensitivity to extrinsic and intrinsic apoptotic stimuli

(Supplementary Fig. 3). Interestingly, however, ablation-triggered and native egress

were not inhibited in these cells even with the addition of 50μM Z-Vad-fmk. Taken

together the events subsequent to chlamydial inclusion rupture and ultimately egress

of the pathogen are not BAK/BAX- or caspase-dependent and is therefore not

classical apoptosis.

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Chlamydial egress is not mediated through a necroptotic Programmed Cell Death

Given the apparently caspase-independent nature of chlamydial inclusion rupture-

induced PCD in what are accepted to be apoptosis-resistant infected cells (Sharma and

Rudel, 2009), we next investigated the likelihood that the mechanism may be

necrotic. The highly responsive and triggerable nature of the death as well as its

dependence upon intracellular calcium suggested that the pathway could be

programmed necrosis or “necroptosis” (Murphy and Silke, 2014). Necroptosis occurs

when cells, in which apoptosis signalling has been blocked, are exposed to death-

inducing stimuli and is mediated through the action of kinase Receptor Interacting

Proteins (RIPs) 1 and 3. RIP1 and RIP3 form supramolecular complexes called the

necrosomes which in turn lead to the activation of the mixed lineage kinase domain-

like protein (MLKL) by RIP3-mediated phosphorylation causing it to homo-

oligomerise (Hildebrand et al., 2014; Wang et al., 2014). Oligomerised MLKL

translocates to the plasma membrane of the cell where it compromises its ability to

preserve intracellular ionic homeostasis. Necroptosis is inhibited by necrostatin-1, a

small molecule inhibitor of RIP1 kinase (Degterev et al., 2008). To determine if the

cell death pathway in question is necroptosis, ablation-triggered and native egress

were examined in the presence of this agent and the rate at which cells died

monitored. Interestingly, treatment with necrostatin-1 did not delay cell death in

either circumstance (Supplementary Fig. 2). To definitively rule out necroptosis,

CRISPr-mediated knockout of RIP1 in HeLa and BAK/BAX double-knockout HeLa

cells was performed (Figure 3D) and both ablation-triggered and native egress

examined (Fig. 3B and 3C). Similar to the observations made earlier, ablation-

triggered and native egress progressed normally in these cells. The egress of the

pathogen is therefore distinct from the established apoptotic and necroptotic PCDs.

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Calpains mediate inclusion rupture

Our laser ablation approach provides us with the unique opportunity to uncouple

inclusion rupture from cell death in a coordinated manner. Intriguingly treatment of

chlamydia infected cells with calpain inhibitors, 100μM calpeptin (Fig 4A) or

PD150606 (data not shown), led to significant persistence of intact chlamydial

inclusions leading to dramatic expansion when compared to control infections.

Eventual inclusion rupture still initiated cell lysis in spite of calpain inhibition (Fig.

4B) as well as ablation-mediated inclusion rupture-induced cell death (Fig. 4C)

indicating that calpain activity was required only for the first stage of chlamydial

egress. Liberation of EBs into the extracellular milieu was measured by conducting

infectious progeny assays using media sampled from cells in the presence and absence

of calpeptin at 72 h p.i. (Fig. 4D & 4E). A marked reduction in extracellular EBs was

observed in calpain-inhibited cells when compared with those treated with the carrier

solvent. Notably, cell lysates treated with calpain inhibitors presented similar numbers

of infectious EBs indicating that calpain inhibition did not adversely impact the

pathogen directly (Fig. 4D & 4E). Therefore, calpains are key regulators of

chlamydial inclusion rupture but not the consequent cell death or the pathogen’s

growth.

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Discussion

In order to propagate an infection, the obligate intracellular pathogen, Chlamydia

trachomatis, must lyse the cell or encompassing extrusion in order to infect new cells.

The resultant cellular damage and inflammation is likely responsible for the localised

scarring and blockage of the fallopian tubes leading to infertility and for the acute

pelvic inflammatory disease frequently associated with infection. Yet, it has long been

recognised that Chlamydia-infected cells are insensitive to both extrinsic and intrinsic

apoptotic stimuli (Fan et al., 1998) and this resistance is requisite for the completion

of the pathogen’s unique dimorphic lifecycle. The resistance to apoptosis appears to

be acquired by a variety of mechanisms including degradation of a key regulator of

apoptosis P53 (Gonzalez et al., 2014; Siegl et al., 2014), but notably, Chlamydia

infected HeLa cells exhibit a marked reduction in caspase 8 enzyme activity (Bohme

et al., 2010). Activated caspase 8 propagates apoptotic signalling by either directly

cleaving and activating downstream effector caspases or by cleaving the BH3 Bcl2-

interacting protein (BID) which translocates to the mitochondria and induces leakage

of cytochrome c. We, however, find no evidence of effector caspase activation

infected cells during native chlamydial egress or following inclusion rupture and

treatment with pan-caspase inhibitors does little to preserve cell or inclusion integrity

of infected cells (Fig. 3A, B &C). We did, however, observe preservation of

neighbouring uninfected cell integrity during chlamydial egress when caspase activity

was inhibited (Fig. 3C and Supplementary Fig. 2B) consistent with in vivo

observations that suggest a role for caspase activity in Chlamydia-induced infertility

(Igietseme et al., 2013). Consistent with this, whilst caspase 1-deficient mice display

much reduced genital tract inflammatory damage following chlamydial infection, the

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mice experienced similar courses of infection indicating that caspase 1 does not play a

direct role in establishment and progression of the infection (Cheng et al., 2008).

How then does Chlamydia kill host cells? In contrast with Perfettini et al (2002), who

observed that BAX translocates to mitochondria in Chlamydia psittaci-infected cells

and that BAX-inhibitor 1 or Bcl-2 expression perturbs C. psittaci-induced caspase-

independent “apoptosis” (Perfettini et al., 2002), here we find that HeLa cells

genome-edited to be profoundly resistant to apoptosis and/or necroptosis through the

selected and iterative deletion of BAK, BAX, and RIP1 as well as in the presence of

pan-caspase and necroptosis inhibitors still die with the same kinetics as control cells

(Fig. 3B & 3C). Whether this disparity with the work of Perfettini et al (2002) reflects

a biological variance unique to the different species of Chlamydia used in the

respective studies remains to be clarified but our data conclusively demonstrates that

the pathway in question is not apoptotic. Recently, RIP3-independent necroptotic

pathways have been described including one that is induced in response to alkylating

DNA-damage agents such as N-methyl-N′-nitro-N′-nitrosoguanidine (MNNG) and

involves the sequential activation of poly(ADP-ribose) polymerase 1 (PARP-1),

calpains, BAX and AIF (Moubarak et al., 2007). Intriguingly, Chlamydia trachomatis

was recently recognised as a DNA damaging agent (Chumduri et al., 2013) and DNA

strand breaks are associated with loss of plasma membrane integrity and organelle

dilation at the later stages of the infection with Chlamydia pneumoniae (Marino et al.,

2008). The chlamydial inclusion rupture-triggered cell death is, however, distinct

from this pathway as HeLa cells do not express RIP3 endogenously (He et al., 2009)

and undergo death independently of BAX (Fig. 3B & 3C).

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We find that Chlamydia-induced cell death is calcium-dependent, occurs regardless of

when in the infection the chlamydial inclusion ruptures and in the presence of

bacterial protein synthesis inhibitors indicating that the lytic process is largely

directed by the host cell (Fig. 1D & 3C). Moreover, our laser-ablation approach

allows us to uncouple inclusion rupture from the subsequent cell death with

unparalleled spatio-temporal resolution and provides the means to examine the action

of specific proteases in the events leading to and following inclusion rupture and cell

lysis without disrupting the pathogen. Notably, application of a small molecule

inhibitor of the chlamydial protease-like activity factor (CPAF) had no impact on

inclusion rupture or cell death and lysis, but served to maintain the integrity of host

cell intermediary filaments as observed by monitoring vimentin-2xGFP in intact live

cells (Supplementary Fig. 1). There has been significant controversy surrounding the

action of this particular chlamydial protein of late owing to a widely published post-

cell lysis in vitro artefact (Chen et al., 2012). Although degradation of vimentin was

clearly refuted, a more recent publication in which CPAF was genetically deleted

from the pathogen provided evidence for a role in post-inclusion rupture disassembly

of vimentin-positive intermediary filaments (Snavely et al., 2014). Our observations

directly support this proposal.

What precisely is released from the inclusion following rupture and therefore triggers

the events that lead to cell death is unknown. Indeed the chlamydial inclusion is a

unique environment which contains an assortment of cellular and bacterial material.

Amongst these are likely a huge array of Pathogen-associated molecular patterns

(PAMPs) and danger signals, many of which could trigger a necrotic response like

that observed in this study. Necrosis is marked by an elevation in ROSs and

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mitochondrial hyperpolarisation but is independent of both caspase activation and

RIP1 (Vanden Berghe et al., 2010): all features observed or consistent with our

observations following chlamydial inclusion rupture. Notably, C. trachomatis L2 does

encode the Chlamydia protein associating with death domains (CADD) (Stenner-

Liewen et al., 2002). Whilst this protein is reported to bind and activate elements of

the host’s apoptotic pathway in vitro, its function has never been examined in vivo. It

would be interesting to monitor the rate of egress and inclusion rupture-triggered cell

death in CADD-null strains should they prove viable.

Finally, whilst inhibition of host caspases had no impact upon inclusion rupture or cell

lysis, inhibition of host cell calpains lead to a dramatic extension in inclusion integrity

without interfering with cell death indicating that calpain-activation is selectively

necessary for chlamydial inclusion lysis. Calpains are a large family of ubiquitous

calcium-sensitive non-lysosomal cysteine proteases responsible for a wide variety of

cellular processes. Our observation is therefore entirely consistent with and extends

upon Hybiske and Stephens’ (2007) original observation that application of a pan-

cysteine protease inhibitor cocktail inhibited inclusion rupture (Hybiske and Stephens,

2007) and is the first reported role for calpains in chlamydial infection. Further

investigation should reveal whether they represent a viable target for therapeutic

intervention during chlamydial infection.

In summary, we have developed a unique laser-ablation method that enables us to

trigger the ordinarily stochastic rupture of the chlamydial inclusion in a controlled and

predictable manner. This has enabled us to segregate the molecular events and

contributions of both the pathogen and the host during chlamydial egress in a

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spatiotemporal manner. Specifically, we demonstrate that the subsequent lytic cell

death pathway observed in Chlamydia infected cells occurs independent of stage of

infection, BAK-, BAX-, RIP1- and caspase-activity and provide further evidence for a

role for the chlamydial effector CPAF in the disassembly of intermediate filaments

following inclusion rupture. Our method highlights the barrier function played by the

inclusion membrane, protecting the encompassed bacteria from cytosolic ROSs, and

thereby highlights the necessity for the pathogen to rapidly escape from the cytoplasm

following inclusion rupture via a non-apoptotic/necroptotic mechanism. Finally we

reveal a novel role for host calpains in the rupture of the chlamydial inclusion but not

the subsequent lytic cell death pathway opening new avenues for investigation.

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Experimental Procedures

Constructs and reagents

CellROX® Green and MitoTracker® Red were supplied by Life Technologies.

Calpeptin, PD150606, Necrostatin-1, carbobenzoxy-valyl-alanyl-aspartyl-[O-methyl]-

fluoromethylketone (Z-Vad-fmk) and VX-765 were supplied by Merck Millipore.

4',6-diamidino-2-phenylindole (DAPI), paraformaldehyde and N-acetyl-L-cysteine

were supplied by Sigma Aldrich. H2B-CFP used was as described previously

(Beronja et al., 2010), Addgene plasmids: 25998). Casper3-GR was supplied by

Evrogen. BAK, BAX and RIP1 polyclonal antibodies were supplied by Cell

Signalling Technology. β-tubulin antibodies were supplied by Li-Cor.

Cell culture, transfection and generation of stable genome-edited and knockout

lines

HeLa cells were maintained in DMEM supplemented with 10% (v/v) FCS and 2mM

L-glutamine (Invitrogen) in a humidified air/atmosphere (5% CO2) at 37oC. Genome-

editing was performed through the sequential generation of stably expressing

mCherry-Cas9 cells followed by the delivery of target specific gRNAs (Aubrey et al.,

2015) in the FH1tUTG lentiviral vector system. BAK gRNA:

GGCCATGCTGGTAGACGTGT. BAX gRNA: TCTGACGGCAACTTCAACTG

(Gong, Huang et al submitted). RIP1 gRNA: AGTGCAGAACTGGACAGCGG.

Clonal populations of cells were isolated by Fluorescence Assisted Cell Sorting.

Chlamydial infection and infectious progeny assay

Chlamydia trachomatis L2 (ATCC VR-902B) and GFP-expressing Chlamydia

trachomatis L2 were generated as described previously (Wang et al., 2011).

20 14/05/y

Chlamydia was used to infect cells at the indicated multiplicity of infection (MOI).

Cells were infected for 2 h in normal DMEM growth media supplemented with 10%

FCS (v/v) and 2mM L-glutamine at 37oC humidified with 5% CO2. After which the

media was replaced with fresh growth media and grown to the stipulated time points.

Infectious progeny assays were performed as previously described (Gonzalez et al.,

2014).

Time-lapse videomicroscopy

For long-term live cell imaging, monolayers were cultured in 35 mm glass‐bottom

dishes (MatTek) or 96 well glass-bottom microplates. Time‐lapse videomicroscopy

was carried out on individual live cells using a Nikon Ti-E inverted deconvolution

microscope using a ×40, 0.9 Plan Apo DIC objective, a Hamamatsu Flash 4.0 4Mp

sCMOS monochrome camera and 37°C incubated chamber with 5% CO2. CFP was

excited with a 438/24nm LED and captured using a 579/40nm emission filter, GFP

was excited with a 485/20nm LED and captured using a 525/30nm emission filter,

mCherry was captured using a 560/25nm LED and captured using a 607/36nm

emission filter.

Laser ablation and confocal time-lapse videomicroscopy

Ablation experiments were performed on an LSM 510 meta Zeiss confocal

microscope at 37°C. Images were acquired using a ×63 objective, 1.4 NA oil Plan

Apochromat immersion lens at ×1.5 digital magnification, with the pinhole adjusted

to 3 Airy units to obtain optical sections 2μm thick. Time-lapse images were acquired

before (3-5 frames) and after ablation with an interval and duration as indicated. A

Ti:sapphire laser (Chameleon Ultra, Coherent Scientific) tuned to 790nm was used to

21 14/05/y

ablate subcellular regions using a constant 12x12 pixel ROI (~0.5-1μm3) was marked

for each experiment and ablated with 1 iteration of a 790nm laser with 65%

transmission. GFP and mCherry fluorescence intensities were monitored before and

after the ablation using a 488nm or 543nm laser for excitation and a 500–550nm or

>560nm emission filters respectively. Donor and Acceptor FRET channels were

acquired using a 488nm laser and emission was collected in the donor emission range

(BP 500-530nm) and acceptor emission range (LP >560nm), respectively.

Correlative Light and Electron Microscopy (CLEM)

For CLEM, cells were seeded and imaged on gridded glass bottom-dishes (MatTek).

Prior to high resolution time-lapse videomicrosopy, low magnification images of the

cells to be examined were captured to obtain their coordinates so that they could be

identified during EM processing. Immediately post-acquisition, cells were fixed in

2.5% glutaraldehyde in PBS and processed for flat embedding in resin (Parton et al.,

1992). After curing, the resin containing the cells was broken away from the plastic

dish. Cells of interest were located by reference to the grid coordinates transferred

onto the resin. 60nm sections were cut parallel to the substratum and imaged after on-

grid staining in a Jeol (Tokyo, Japan) 1011 transmission electron microscope.

Western Immunoblotting

Infected and uninfected cell monolayers were lysed directly with SDS lysis buffer

(100 mM Tris/HCL, pH 6.8, 4% SDS, 20% glycerol, 0.02% bromophenol blue, 200

nM dithhiothreitol) and immediately boiled at 95oC for 10 min. Equal amounts of

protein were loaded, resolved on 10% SDS-polyacrylamide gels and transferred onto

Immobilon-FL PVDF membranes (Millipore, USA) according to the manufacturer’s

22 14/05/y

instructions. Western immunoblotting using ECL was performed as described

previously (Yang et al., 2015).

Apoptosis and Cell Survival Assays

Extrinsic and intrinsic apoptosis was induced through the application of 50ng/ml

recombinant human TNFα (ORF Genetics) and 10 µg/ml cycloheximide (Sigma

Aldrich) or 2 µg/ml staurosporine (Sigma Aldrich). Cells were incubated for 24 h

until fixation with 4% paraformaldehyde in phosphate buffered saline (PBS). Cell

viability was determined by monitoring the presence of soluble mCherry-Cas9 using a

Nikon Ti-E inverted deconvolution microscope using a ×40, 0.9 Plan Apo DIC

objective, a Hamamatsu Flash 4.0 4Mp sCMOS monochrome camera. Single cell live

imaging of caspase 3 activity was conducted using the caspase 3 apoptosis sensor

Casper3-GR as per manufacturer’s instructions (Evrogen).

23 14/05/y

Author Contributions

MCK designed and conducted all of the experiments, wrote the manuscript and

constructed the figures. GG and ASY assisted in the development and optimisation of

the laser-ablation protocol employed throughout the manuscript and provided useful

input into the manuscript. CF and RP conducted the correlative light and electron

microscopy presented in Figure 2. DS, MCT and JMM developed, optimised and

supplied the CRISPr and gRNA technology presented in Figure 3. WM provided

chlamydia strains and useful input into the manuscript. RDT designed experiments

and wrote the manuscript.

24 14/05/y

Acknowledgments

This work was supported by funding from the National Health and Medical Research

Council (NHMRC) of Australia (606788) and the Australian Research Council

(DP150100364). MCK was supported by an Australian Research Council Discovering

Early Career Researcher Award (DE120102321). MCT was supported by a Victorian

International Research Scholarship. The NHMRC provided fellowship support to

JMM (APP1105754), ASY (APP1044041), RGP (APP569542), and RDT

(APP1041929). RGP and ASY also acknowledge support from NHMRC Program

Grant (APP1037320). DS, MCT and JMM acknowledge support from NHMRC

IRIISS (grant 9000220) and Victorian Government Operational Infrastructure Support

schemes. The authors acknowledge the facilities, and the scientific and technical

assistance, of the Australian Microscopy & Microanalysis Research Facility at the

Centre for Microscopy and Microanalysis, The University of Queensland, and the

Australian Cancer Research Foundation (ACRF)/Institute for Molecular Bioscience

(IMB) Dynamic Imaging Facility for Cancer Biology. Finally, we thank David Segal,

David Huang, John Silke and Marco Herold (he Walter and Eliza Hall Institute of

Medical Research, Australia) for reagents and helpful discussions in the preparation

of this manuscript.

25 14/05/y

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Figure Legends:

Figure 1: Laser-mediated inclusion rupture triggers chlamydial egress. A) Time-

lapse videomicroscopy of GFP-expressing mCherry-Rab25 and CFP-H2B stable

HeLa cells imaged from 36h p.i. with CTL2 (MOI~0.5). Presented is a single egress

even captured from 49 h p.i. Arrows highlight the nuclei and asterisks the inclusion.

Interval of capture was 5 min. B) Diagram of inclusion ablation system and time-lapse

videomicroscopy of the system applied on mCherry-Rab25 stable HeLa cells 48 h p.i.

with GFPCTL2 (MOI~0.5). Arrows highlight the nuclei, asterisks the inclusion and

arrow heads blebbing of the plasma membrane. Interval of capture is as indicated. C)

Time-lapse videomicroscopy of mCherry-Rab25 stable HeLa cells ablated 24h p.i.

with GFPCTL2 (MOI~0.5). Arrows highlight the nuclei and arrow heads blebbing of

the plasma membrane. Interval of capture is as indicated. D) Quantification of cell

survival of ablated and unablated cells under the conditions as indicated. >10 cells

were quantified for each condition. E) Time-lapse videomicroscopy of mCherry-

Rab25 stable HeLa cells ablated either 24h p.i. with GFPCTL2 (MOI~0.5) in

calcium-free Ringer’s solution and 50μM BAPTA-AM. Interval of capture is as

indicated. F) Time-lapse videomicroscopy of mCherry-Rab25 stable HeLa cells

ablated 24h p.i. with GFPCTL2 in the presence of 800nM YM201636 to swell

endosomal compartments. Insets highlight a ruptured Rab25-positive chlamydial

inclusion denoted by the prominent GFP-expressing bacterial fluorescence within and

a ruptured Rab25-positive endosome. Arrows highlight the nuclei, asterisks the

inclusion and arrow heads blebbing of the plasma membrane.

Figure 2: Correlative light and electron microscopy reveals the barrier function of

the chlamydial inclusion. A) Time-lapse videomicroscopy of GFP-Rab25 stable HeLa

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cells ablated 24h p.i. with CTL2 (MOI~0.5). B) Transmission electron micrographs of

these same cells. Arrows highlight swollen chlamydial RBs. C) Time-lapse

videomicroscopy of mCherry-Rab25 stable HeLa cells ablated 24h p.i. with CTL2

(MOI~0.5) in the presence of CellROX® Green. D) Quantification of intra-

inclusional CellROX® Green fluorescence. N= 5, Error bars present the Standard

Deviation from the Mean. E) Time-lapse videomicroscopy of GFP-Rab25 expressing

stable HeLa cells ablated 24h p.i. with CTL2 (MOI~0.5) in the presence of

Mitotracker® Red CMXRos. Arrows highlight swelling mitochondria and asterisks

the inclusion.

Figure 3: Chlamydia-induced cell death is not apoptotic or necroptotic A) Time-lapse

videomicroscopy of Casper3-GR expressing HeLa cells ablated 24h p.i. with CTL2

(MOI~0.5). B) Quantification of inclusion-rupture induced cell death under the

indicated conditions. C) Quantification of native egress-induced cell death under the

indicated conditions. Presented in pie-charts are the proportions of primary infected,

≥secondary infected and uninfected cells surviving cells at the indicated time-point as

monitored by GFP-CTL2 fluorescence. N=3, Error bars present the Standard

Deviation from the Mean. For clarity, only error bars for DMSO and

chloramphenicol are presented. D) Western immunoblotting of HeLa cells genome-

edited for the indicated targets.

Figure 4: Calpains mediate inclusion rupture. A) Time-lapse videomicroscopy of

mCherry-Rab25 stable HeLa cells infected from 8-72 h p.i. with GFP-CTL2

(MOI~0.5) in the presence or absence of 100M calpeptin. B) Quantification of native

egress-induced cell death under the indicated conditions. Presented in pie-charts are

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the proportions of primary infected, ≥secondary infected and uninfected cells

surviving cells at the indicated time-point as monitored by GFP-CTL2 fluorescence.

N= 3, Error bars present the Standard Deviation from the Mean. C) Quantification of

inclusion-rupture induced cell death under the indicated conditions. D) mCherry-

Rab25 and CFP-H2B stable HeLa cells infected with the indicated diluents from

HeLa cells infected for 72 h p.i. with GFP-CTL2 (MOI~0.5). Presented images were

captured 24 h p.i.. E) Infectious progeny forming units in media and whole cell

lysates from infected cells cultured in the presence or absence of calpeptin were

quantified. . N= 3, Error bars present the Standard Deviation from the Mean.

Supplementary Figure 1:

Time-lapse videomicroscopy of 2xGFP-vimentin-expressing mCherry-Rab25 stable

HeLa cells infected for 24 h p.i. with CTL2 (MOI~0.5) and ablated in the presence

and absence of a CPAF-inhibitory peptide. Arrows highlight the nuclei and asterisks

the inclusion.

Supplementary Figure 2:

A) Quantification of inclusion-rupture induced cell death under the indicated

conditions. B) Quantification of native egress-induced cell death under the indicated

conditions. Presented in pie-charts are the proportions of primary infected,

≥secondary infected and uninfected cells surviving cells at the indicated time-point as

monitored by GFP-CTL2 fluorescence. N=3, Error bars present the Standard

Deviation from the Mean. For clarity, only error bars for DMSO and VX765 are

presented.

34 14/05/y

Supplementary Figure 3:

Live images of genome-edited HeLa cells following extrinsic and intrinsic apoptotic

stimuli at the indicated time-points.

35 14/05/y

Movie Legends:

Supplementary Movie 1:

Time-lapse videomicroscopy mCherry-Rab25 and CFP-H2β stable HeLa cells

infected for 48 h p.i. with CTL2 (MOI~0.5) as presented in Figure 1A. Interval is as

indicated.

Supplementary Movie 2:

Time-lapse videomicroscopy mCherry-Rab25 and GFP stable HeLa cells infected for

48 h p.i. with CTL2 (MOI~0.5) ) as presented in Figure 1B. Inclusion was ablated as

indicated (reticule). Interval is as indicated.

Supplementary Movie 3:

Time-lapse videomicroscopy mCherry-Rab25 stable HeLa cells infected for 24 h p.i.

with GFPCTL2 (MOI~0.5) ) as presented in Figure 1C. Inclusion was ablated as

indicated (reticule). Interval is as indicated.

Supplementary Movie 4

Time-lapse videomicroscopy GFP-Rab25 stable HeLa cells infected for 24 h p.i. with

CTL2 (MOI~0.5) and transferred to Calcium-free Ringer’s solution containing 50M

BAPTA-AM immediately prior to imaging as presented in Figure 1E. Inclusions were

ablated as indicated (reticule). Interval is as indicated.

Supplementary Movie 5

Time-lapse videomicroscopy GFP-Rab25 stable HeLa cells infected for 24 h p.i. with

CTL2 (MOI~0.5) and transferred to media containing 800nM YM801636

36 14/05/y

immediately prior to imaging as presented in Figure 1F. Inclusions were ablated as

indicated (reticule). Interval is as indicated.

Supplementary Movie 6

Time-lapse videomicroscopy mCherry-Rab25 stable HeLa cells infected for 24 h p.i.

with CTL2 (MOI~0.5) in the presence of CellROX® Green. Inclusions were ablated

as indicated (reticule). Interval is as indicated.

Supplementary Movie 7

Time-lapse videomicroscopy GFP-Rab25 stable HeLa cells infected for 24 h p.i. with

CTL2 (MOI~0.5) in the presence of Mitotracker® Red CMXRos. Inclusions were

ablated as indicated (reticule). Interval is as indicated.

Supplementary Movie 8

Time-lapse videomicroscopy mCherry-Cas9 stable HeLa cells treated with DMSO

imaged from 35 h p.i. with CTL2 (MOI~0.5) as presented in Figure 4A. Interval is as

indicated.

Supplementary Movie 9

Time-lapse videomicroscopy mCherry-Cas9 stable HeLa cells treated with calpeptin

imaged from 35 h p.i. with CTL2 (MOI~0.5) as presented in Figure 4A. Interval is as

indicated.

37 14/05/y