ABSTRACT
Our understanding of how the obligate intracellular bacterium Chlamydia trachomatis reprograms the cell biology of host cells in the upper genital tract is largely based on observations made in cell culture with transformed epithelial cell lines. Here we describe a primary spherical organoid system derived from endometrial tissue to recapitulate epithelial cell diversity, polarity, and ensuing responses to Chlamydia infection. Using high-resolution and time-lapse microscopy, we catalogue the infection process in organoids from invasion to egress, including the reorganization of the cytoskeleton and positioning of intracellular organelles. We show this model is amenable to screening C. trachomatis mutants for defects in the fusion of pathogenic vacuoles, the recruitment of intracellular organelles, and inhibition of cell death. Moreover, we reconstructed a primary immune cell response by co-culturing infected organoids with neutrophils, and determined that the effector TepP limits the recruitment of neutrophils to infected organoids. Collectively, our model details a system to study the cell biology of Chlamydia infections in three dimensional structures that better reflect the diversity of cell types and polarity encountered by Chlamydia upon infection of their animal hosts.
Summary statement 3D endometrial organoids to model Chlamydia infection and the role of secreted virulence factors in reprogramming host epithelial cells and immune cell recruitment
INTRODUCTION
Chlamydia trachomatis is a clinically important pathogen, responsible for the majority of sexually transmitted bacterial infections (WHO, 2018). In the female upper genital tract (UGT), chronic and untreated asymptomatic infections can lead to pelvic inflammatory disease, tubal scarring and infertility (Haggerty et al., 2010), and are associated with cancers of the cervix and endometrium (Koskela et al., 2000). Yet, despite decades of studies the molecular mechanisms by which C. trachomatis disrupts epithelial tissue structure and induces pathology in the UGT are not well understood.
The genus Chlamydia is comprised of nine Chlamydia species that infect eleven different vertebrate hosts, including the human pathogen C. trachomatis and rodent-adapted C. muridarum (Horn, 2008; Schachter, 1978). Based on sequence similarity and disease manifestations, C. trachomatis is further classified into serovars and biovars that target the genital tract (serovars D-K) and ocular epithelia (serovars A-C), the etiological cause of the inflammatory disease trachoma (Stephens et al., 2009). The lymphogranuloma venereum (LGV) biovar (serovars L1-L3) is an invasive infection in the urogenital or anorectal tracts that disseminates to the lymph nodes (Mabey and Peeling, 2002). Of all the serovars, Chlamydia LGV serovar L2 is the only one that is fully tractable to molecular genetic manipulations (Mueller et al., 2016; Wang et al., 2011).
All Chlamydia transition between two developmental forms: the environmentally stable and infectious elementary body (EB), and the non-infectious but replication-competent reticulate body (RBs) (Abdelrahman and Belland, 2005). To invade cells, EBs use a type III secretion system (T3S) to deliver effector proteins directly into host epithelial cells. These invasion-associated early T3S effectors stimulate rearrangements in filamentous actin (F-actin) at attachment sites to promote entry and uptake into an intracellular membrane vacuole termed the “inclusion.” (Elwell et al., 2016). Additional effectors are then secreted and inserted into the nascent inclusion membrane. These effectors, termed inclusion membrane (Inc) proteins, co-opt microtubule-based trafficking to aid in the migration of inclusion to the microtubule organizing center, subvert intracellular organelles, including the Golgi apparatus, endoplasmic reticulum, peroxisomes, and lipid droplets, and establish membrane contact sites to presumably facilitate the acquisition of nutrients form its host (Agaisse and Derré, 2014; Boncompain et al., 2014; Derré et al., 2011; Dumoux et al., 2012; Grieshaber et al., 2003; Kumar et al., 2006; Mital et al., 2015; Moore et al., 2008). Within the inclusion, EBs differentiate into RB forms, replicate and expand along the perimeter of the inclusion. The inclusion itself is wrapped in a network of actin and intermediate filaments, microtubules, and septins (Dumoux et al., 2015; Kumar and Valdivia, 2008; Volceanov et al., 2014; Wesolowski et al., 2017). RBs asynchronously differentiate back into EBs and escape from the cell through a F-actin-mediated extrusion of the inclusion or host cell lysis to initiate the next round of infections (Hybiske and Stephens, 2007).
Chlamydia maintains an intracellular replicative niche by suppressing cell-autonomous innate immunity (Finethy and Coers, 2016). For example, Chlamydia-driven rearrangements of the cytoskeleton promotes inclusion stability, limiting its detection and activation of the innate immune response (Kumar and Valdivia, 2008). Chlamydia Inc proteins counteract host defenses by deubiquitinating proteins at the inclusion, inhibiting the lytic activity of interferon induced factors (e.g. guanylate binding proteins) and suppressing cell death by subverting membrane trafficking to inhibit host surveillance programs and apoptosis (Coers et al., 2008; Faris et al., 2019; Fischer et al., 2017; Sixt et al., 2017). Despite these activities, Chlamydia is still recognized by the host cell which can lead to a pro-inflammatory response (Rasmussen et al., 1997), including the expression and secretion of chemokines, such as the neutrophil attractants interleukin-6 (IL-6) and interleukin-8 (IL-8) (Buchholz and Stephens, 2008; Rasmussen et al., 1997).
Our understanding of the cell biology of how Chlamydia re-programs host cellular functions is based largely on studies in two-dimensional cell culture systems with transformed cervical epithelial cell lines like HeLa cells. While these models are experimentally tractable and have been of great use, they do not accurately represent the natural sites of infection: polarized UGT epithelial cells that have a distinct spatial organization of the cytoskeleton, organelle positioning, and signaling modules that are distinct from transformed nonpolarized epithelia (Dolat and Valdivia, 2019; Rodriguez-Boulan and Macara, 2014). Infections in polarized epithelia have uncovered unique aspects of the interaction between Chlamydia and its target cell: C. trachomatis replication is enhanced in polarized epithelial cells and the inclusion preferentially intercepts vesicles destined for the basolateral side of infected cells (Guseva et al., 2007; Moore et al., 2008). Infections in primary polarized human ecto- and endocervical epithelial explants revealed that Chlamydia alters epithelial structure by inducing epithelial-to-mesenchymal transition (Zadora et al., 2019). Furthermore, long-term Chlamydia infections in fallopian tube organoids, self-renewing primary three-dimensional epithelial tissue-like cultures, promotes epithelial stemness, cellular proliferation, and modifications to genome organization often associated with ageing tissues (Kessler et al., 2019). These observations may help explain some of the associations observed between Chlamydia infections and cervical and ovarian cancers (Shanmughapriya et al., 2012; Zhu et al., 2016).
Three-dimensional organoids have emerged as a compelling ex-vivo system to study epithelial cell biology as they better recapitulate aspects of the cellular diversity, structure and function of epithelial tissues (Rossi et al., 2018). Isolated human and mouse glandular endometrial epithelia cultured in a chemically defined media were recently reported to form endometrial organoids (EMOs) comprised of hormonally responsive polarized epithelial subtypes (e.g. ciliated, secretory) that mimic aspects of the estrous cycle and implantation (Boretto et al., 2017; Turco et al., 2017). EMOs exhibit long-term genetic stability and are amenable to high-throughput screens (Boretto et al., 2019). Here, we describe an infection model that uses EMOs and microinjection to deliver Chlamydia to the apical surface of polarized epithelium, thus mimicking the natural route of infection. Using both the rodent-adapted C. muridarum and the genetically-tractable human pathogen C. trachomatis serovar L2, we demonstrate key aspects of the infection cycle and cellular activities driven by C. trachomatis effectors. We further show that we can recapitulate aspects of cellular immunity in infected EMOs by monitoring the recruitment and behavior of neutrophils. Using a C. trachomatis T3S effector mutant that alters immunity-related signaling pathways, including the expression of neutrophil chemoattractants, we further demonstrate that this system can be used to measure the manipulation of host immune responses by Chlamydia virulence factors.
RESULTS
Cells harvested from the upper genital tract develop into endometrial organoids of cellular composition similar to that of primary tissues
Isolated endometrial glandular epithelia were cultured in a three-dimensional Matrigel matrix in the presence of conditioned medium from L-WRN cells. L-WRN cells stably express and secrete Wnt-3A, R-Spondin 3, and Noggin, efficiently generate organoids from various mouse tissues with little batch-to-batch variation (Miyoshi and Stappenbeck, 2013; VanDussen et al., 2019). Indeed, EMOs readily form over the course of one week and grow into large, spherical structures (Fig 1 B-C).
Using high-resolution confocal microscopy, we determined that EMOs are comprised of a single layer of polarized epithelial cells surrounding a large, hollow lumen. The epithelia express and secrete mucin (Muc1) (Fig 1D) and establish adherens and tight junctions (Fig 1E-F), respectively marked by E-cadherin and zonula occludens-1 (ZO-1). To assess the integrity of the spheroids and the functionality of the cell-cell junctions, we microinjected fluorescent 3 kDa dextran into the organoid lumen. The fluorescent tracers remained in the lumen indicating that barrier function is maintained over prolonged periods of time (Fig 1G). Endometrial tissues are responsive to sex hormones, increasing cellular proliferation and altering gene expression and cell type abundance in the presence of estrogen and progesterone. EMOs are similarly responsive as assessed by the larger increase in size upon exposure to estrogen for four days, as compared to untreated EMOs (2.0x vs 1.37x), and the formation of multi-ciliated epithelia as marked by acetylated microtubules (Fig 1H-I). Overall, the method produces hormonally responsive polarized EMOs with intact barrier function and a distinct lumen.
Chlamydia infection of endometrial epithelial leads to cytoskeletal reorganization and disruption of cell-cell junctions
To mimic the natural route of Chlamydia invasion we microinjected GFP-expressing C. trachomatis serovar L2 or C. muridarum CM006, a high pathology isolate (Poston et al., 2018), into the EMO lumen. We monitored the subcellular localization of F-actin and β-catenin, which are known to localize to the inclusion (Kessler et al., 2012) at early timepoints (8-16 hours) post injection. EBs readily invaded the polarized epithelial cells, promoted the assembly of actin filaments as the nascent inclusion formed, and recruited β-catenin (Fig 2A-B). Consistent with a previous ex vivo fallopian tube infection models (Kessler et al., 2012), infected epithelia showed a striking loss of apical F-actin and cell-cell junction integrity, demarcated by more diffusive β-catenin localization, indicating that epithelial polarity and barrier function are compromised during invasion. We observed β -catenin recruitment to inclusions only at very early time points with no significant recruitment to inclusions by 48 hpi (Fig S1A-B). More recently, Chlamydia has been shown to target epithelial tight junctions, altering the expression of tight junction proteins and reducing transepithelial electrical resistance (Kumar et al., 2019). Using EMOs derived from ZO1-GFP expressing transgenic mice (Foote et al., 2013), we show that ZO-1 is recruited to EBs during invasion along with β-catenin, which may further underlie the disruption to epithelia barrier functions (Fig 2C). Taken together, these data show that Chlamydia invasion remodels the F-actin network and cell-cell junction organization in a manner that disrupts cell polarity early during the infectious process.
During the mid and late stages of infection Chlamydia remodels the cytoskeleton of HeLa cells at the periphery of inclusions (Kumar and Valdivia, 2008). We monitored the localization of various cytoskeletal elements in infected EMOs at mid-cycle (24 hpi) by probing for F-actin and immunostaining against microtubules, cytokeratins, and septins – all of which localize to the inclusion periphery (Chin et al., 2012; Kumar and Valdivia, 2008; Volceanov et al., 2014). In polarized epithelial the cytoskeletal architecture is markedly different from non-polarized and transformed cells. For example, the F-actin network is enriched at the apical membrane, microvilli, and cell-cell junctions rather than a dense network of bundled actin stress fibers (Fig 2A). Moreover, in contrast to 2D culture settings, microtubule organizing centers localize proximal to the apical membrane and generate microtubules that extend towards the basolateral domain, often along the lateral membrane (Pickett et al., 2019). We observed F-actin structures around the inclusion, either as filamentous-like rings or discrete puncta, which may depend on the stage of the infection cycle (Fig 2E). Moreover, F-actin signal on the apical membrane was markedly reduced or absent in infected epithelia compared to uninfected neighboring cells (Fig 2F), suggesting that infection and/or inclusion growth promotes the loss of microvilli. In uninfected EMOs, microtubules run along the apico-basolateral axis and terminate at the basolateral membrane (Fig 2G). During infection, we observed microtubules and acetylated microtubules prominently assembled around the inclusion (Fig 2H-I). Because Chlamydia effectors, such as CT288 and IPAM, target centrosomal proteins (Almeida et al., 2018; Dumoux et al., 2015), this model can be used to explore effector functions in MT assembly during infection.
The organization and dynamics of F-actin and microtubules are regulated directly by septins, a conserved family of GTPases that assemble into heteropolymers and filaments (Mostowy and Cossart, 2012; Spiliotis, 2018). In EMOs, septin 2 (Sept2) localizes to the apical and lateral membranes and partially co-localizes with F-actin at the basolateral surface (Fig S1C). In infected EMOs Sept2 undergoes a dramatic reorganization to forms filamentous rings around the inclusion (Fig S1D) as had been previously observed in non-polarized cells (Volceanov et al., 2014).
Finally, we probed the endometrial epithelia for intermediate filaments. Both keratins and vimentin localize around the inclusion in transformed epithelia and their absence disrupts inclusion stability (Kumar and Valdivia, 2008; Tarbet et al., 2018). As expected, EMO epithelia express keratins (Fig S1E) which relocalize to the C. trachomatis inclusion (Fig S1F). However, unlike transformed epithelia, EMO epithelial cells do not express vimentin (Fig S1G), and its expression is not induced during infection with C. muridarum (Fig S1H). In contrast, primary endometrial stromal fibroblasts infected with C. muridarum and C. trachomatis displayed prominent vimentin filaments cages surrounding inclusions (Fig S1I).
Overall these findings indicate that despite the difference in cytoskeletal organization between transformed, non-polarized epithelial cells, Chlamydia likely employs conserved mechanism to reorganize these structures during cell entry and formation of inclusions.
Live imaging of inclusion expansion, dynamics and exit
We developed an imaging platform to monitor inclusion growth, dynamics and exit using timelapse 3D spinning disk confocal microscopy. EMOs were infected for 24 hours with C. trachomatis serovar L2 expressing GFP and subsequently imaged live for 16-18 hours. We noted inclusion expansion and dynamics, including the apparent fusion of adjacent inclusions (Fig 3A). We next tested if inclusion fusion is a phenomena that occurs in EMOs by co-infecting with C. trachomatis L2 expressing GFP or mCherry, or with a C. trachomatis mutant (M923) lacking the fusogenic factor IncA (Kokes et al., 2015; Sixt et al., 2017). GFP and mCherry positive inclusions were readily apparent EMOs co-infected with wild-type strains (Fig 3B) but not in EMOs coinfected with the IncA mutants (Fig 3B), indicating that this model can be employed to investigate the propensity of inclusions to fuse and if this process is influenced by the cell type being infected.
We next monitored inclusions exit dynamics using EMOs derived from ROSAmTmG mice that express a membrane-targeted tdTomato allowing for visualization of the plasma membrane of individual cells. First, EMOs were infected with either GFP-expressing C. muridarum for 24 hours or C. trachomatis L2 for 48h and imaged live for an additional 16 hours. We observed that C. muridarum inclusions were released exclusively into the organoid lumen via lysis or extrusion, which left the infected cell intact, suggesting that apical exit may be the natural mechanism by which C. muridarum escapes the host cell (Fig 3 D-E). In contrast, C. trachomatis L2 inclusions were observed to extrude both apically into the lumen (Fig. 3F) and basolaterally into the extracellular matrix (Fig. 3G).
The reorganization of the Golgi apparatus in infected endometrial epithelia requires the inclusion membrane protein InaC
Host cell organelles are recruited to Chlamydia inclusions to form tethering complexes and to intercept lipid-rich vesicles and acquire nutrients (Elwell and Engel, 2012). The inclusion membrane proteins IncD, InaC, and Cdu1 have been determined to target Golgi-resident proteins and potentially regulate/intercept polarized trafficking (Agaisse and Derré, 2014; Derré et al., 2011; Kokes et al., 2015; Pruneda et al., 2018). During epithelial polarization, the Golgi apparatus migrates and expands above the nucleus, proximal to the apical membrane, where it regulates the polarized sorting of vesicles and proteins to distinct membrane domains (Bacallao et al., 1989). Using high-resolution confocal microscopy, we observed that the Golgi apparatus is more widely dispersed in EMO epithelial cells than in HeLa cells, often surrounding the nucleus and extending toward the apical surface (Fig. 4A). Consistent with previous studies (Heuer et al., 2009; Kokes et al., 2015; Pokrovskaya et al., 2012; Pruneda et al., 2018; Rejman Lipinski et al., 2009), however, we found that during early to mid-infection inclusions recruit and reorganize the Golgi apparatus (Fig. 4B).
We next visualized the extent to which the Golgi apparatus is re-positioned around the C. muridarum and C. trachomatis inclusions. Indeed, both species recruit the Golgi around the inclusion periphery to a similar extent (Fig. 4C). Because the recruitment of Golgi stacks to the inclusion is regulated by the expression and secretion of Inc proteins, such as InaC, we sought to determine if this phenotype is recapitulated in InaC mutants. EMOs were infected with the C. trachomatis mutant M407, bearing an InaC truncation (InaCQ103*), M407 complemented with inaC or an empty vector control (Kokes et al., 2015). Indeed, the InaC-deficient C. trachomatis failed to promote the redistribution of Golgi around the inclusion of infected EMO cells, while the complemented mutant shows similar distribution to that of the wild-type infected EMO cells (Fig. 4D). Collectively, these data show that despite the difference of Golgi complex morphology in polarized EMO epithelia, this organelle is still repositioned at the inclusion periphery and that this process is mediated by the same effectors.
CpoS provides protection from Chlamydia-induced cell death in endometrial epithelia
Pathogen-mediated remodeling of organelle dynamics and organization can promote infection by blocking cell-autonomous immunity. For example, host recognition of microbial DNA can induce the translocation of STING (stimulator of interferon genes) from the ER to the Golgi, where it activates the interferon response (Ishikawa et al., 2009). The type I interferon response in Chlamydia-infected cells requires STING (Barker et al., 2013; Prantner et al., 2010), and the Chlamydia Inc CpoS functions to dampen the interferon response by blocking STING translocation and cell death (Sixt et al., 2017). CpoS is among a subset of Incs recently identified to promote inclusion integrity and the viability of infected cells (Sixt et al., 2017; Weber et al., 2017).
To monitor the degree of cell death in infected EMOs, we microinjected EMOs with C. trachomatis L2 in the presence of propidium iodide (PI) and imaged live at 24 hpi. In both uninfected and infected EMOs we observed little to no PI staining, indicating that Chlamydia efficiently blocks the induction of any cell death mediated defense mechanisms (Fig. 5A). We next tested if C. trachomatis deficient in the pro-survival factor CpoS would lead to cell death by infecting EMOs with the C. trachomatis mutant M007, encoding a truncated CpoSQ31*. Indeed, endometrial epithelial cells containing CpoS-deficient inclusions were frequently positive for PI, indicating the activation of cell death pathways (Fig. 5B). We measured the extent of this phenotype by quantifying PI signal frequency in each infected EMO and show a consistent increase with the CpoS-deficient strain (Fig. 5B-C). These observations highlight that the molecular mechanisms used by Chlamydia to protect infected cells from cell death-based antimicrobial responses in conserved in endometrial epithelial cells.
Reconstitution of immune cell recruitment to Chlamydia infected EMOs
Chlamydia promotes the recruitment of immune cells, including neutrophils, which interact with and can invade infected epithelia to make direct contact with inclusions in infected animals (Rank et al., 2011). Their recruitment and activation at infection foci can damage the UGT (Lacy et al., 2011; Lee et al., 2010a; Lijek et al., 2018). However, Chlamydia has evolved strategies to limit the function of neutrophils. For example, the Chlamydia protease CPAF can inhibit neutrophil activation and netosis by cleaving neutrophil receptors (Rajeeve et al., 2018). Thus, we explored the application of the infection model by reconstituting aspects of the engagement of cellular immunity and its potential impact on Chlamydia infection. We co-cultured infected EMOs with primary bone marrow derived neutrophils and used high-resolution and time-lapse microscopy to quantify and visualize their recruitment to EMOs and interactions with infected epithelial cells.
EMOs were microinjected with mCherry-expressing C. trachomatis L2 or fluorescently labeled dextran. At 4 or 20 hours post-infection, primary fluorescently labelled neutrophils (CellTracker™) were added to the media for an additional 20 hours prior to imaging (Fig. 6A). Neutrophils infiltrated the Matrigel and migrated specifically to infected EMOs (Fig. 6A-B). EMOs microinjected with fluorescent dextran failed to recruit a significant number of neutrophils, indicating that the transient damage from the microinjection does not induce a chemoattracting response (Fig. 6B). We observed a subset of neutrophils make contact directly with infected EMO epithelial cells and even inclusions but did not appear to undergo netosis (Fig. 6C-D). Using timelapse spinning disk microscopy, we visualized neutrophil dynamics at infected EMOs and again observed a subset of neutrophils interact with the infected EMO and make contact with an intracellular inclusion (Fig. 6E-F).
Infected epithelial cells secrete IL-6, IL-8, and members of the CXC chemokine family that could drive the recruitment of neutrophils towards Chlamydia-infected cells (Dessus-Babus et al., 2000; Frazer et al., 2011; Rasmussen et al., 1997). In mice, the absence of CXCR2, the CXC chemokine receptor, reduces acute inflammation and pathology in the UGT without affecting bacterial burden (Lee et al., 2010b). A transcriptional analysis of infected endocervical epithelial cells, indicated that the Chlamydia effector TepP enhances early type I interferon responses and dampens the expression of interleukin-6 (IL-6) and CXCL3 (Chen et al., 2014), chemokines that promote neutrophil chemotaxis (Fielding et al., 2008; Wright et al., 2014). We tested if TepP played a role in in regulating neutrophil infiltration by infecting EMOs by first generating a TepP knock out mutant by targeted insertion of an aadA cassette into the tepP locus (Fig. 6G). This mutant was then transformed with complementing plasmid expressing TepP under its native promote (+ tepP) or an empty vector (− tepP), and the resulting strains microinjected into EMOs followed by co-culturing with primary neutrophils at 4 hours post microinjection. Infected EMOS were imaged live for 20 hours. C. trachomatis strains lacking TepP showed significantly more neutrophil recruitment to the EMO, suggesting that TepP acts to dampen the innate immune response by limiting neutrophil influx (Fig. 6H-I).
Collectively, these data show that we can reconstitute the infiltration of innate immune cells to infected epithelia and address the role of Chlamydia virulence factors regulating the immune response.
DISCUSSION
We developed a primary organoid model system to interrogate the cell biology Chlamydia infections in a cellular context that better mimics the architecture and diversity of cells present in the UGT epithelia, the target of these pathogens. Despite differences in the cytoskeletal and endomembrane organization in EMO cells compared to traditional two-dimensional tissue culture systems, we found remarkable conservation in the type of intracellular processes targeted by Chlamydia and in the effector proteins used. Our observations complement other recently reported models, including a fallopian tube organoid infection model (Kessler et al., 2019), and an endometrial organoid infection model that shows Chlamydia replicates within EMO epithelia and is amenable to inducible expression of Chlamydia Incs (Bishop et al. 2020, In Press). These models, however, do not infect the apical surfaces of intact organoids, which precludes the ability to monitor the natural route of Chlamydia invasion.
We provide evidence that Chlamydia reprograms the epithelial cytoskeleton and disrupts epithelial structure, at least transiently, during apical invasion and inclusion formation. These results are in agreement with infections in fallopian tube explants where Chlamydia infection disrupts epithelial architecture, altering the organization of polarity proteins in infected and uninfected neighboring cells (Kessler et al., 2012). Although we observe marked recruitment of β-catenin to early inclusions and Chlamydia entry sites, this localization does not occur in mid-to-late stage inclusions. Nevertheless, β-catenin promotes Chlamydia infection, possibly through Wnt-based signaling as pharmacological inhibition of Wnt signaling in transformed endometrial epithelial cells reduces Chlamydia growth (Kintner et al., 2017).
An advantage of this model lies in the ability to generate organoids from transgenic mouse lines. For instance, we used the ROSAmTmG to better delineate single cell boundaries while following the dynamics of inclusions exiting the host cell. While all of the C. muridarum inclusions we observed exited apically, we observed a subset of C. trachomatis L2 inclusions positioned towards and exiting through the basolateral membrane, reminiscent of the observation in polarized endometrial cancer cells infected with C. trachomatis serovar E and treated with tamoxifen (Hall et al., 2011). It is unclear whether the exit strategy depends on the organization of polarized epithelial cell, cell-type specific responses, or serovar-specific functions. Because EMOs are comprised of a multiple epithelial subtypes (Fitzgerald et al., 2019), this model can be used to identify cell type-specific responses to infection.
We extended the application of EMOs to monitor the interaction between primary immune cells and infected epithelial cells. The rapid and specific recruitment of neutrophils to infected EMOs indicate that this model will be useful to dissect the dynamics of immune cells interaction with Chlamydia infected cells and the role played by host factors that regulate the innate immune responses. Neutrophils appear to have a limited role in clearing the infection, but rather influence the adaptive immune response by promoting the recruitment of T cells (Lacy et al., 2011). Studies using a combination of host and Chlamydia genetics can further identify the interaction between Chlamydia and the innate and adaptive immune response. For instance, we tested the effect of the T3S effector TepP on the influx of neutrophils to infected EMOs. Because neutrophil infiltration is regulated by type I and type III interferons (Blazek et al., 2015; Seo et al., 2011), TepP-dependent regulation of the interferon response and chemokine expression may function in part to reduce immune cell infiltrates and promote survival in the UGT.
In conclusion, we provide a high-resolution blueprint for the cell biology of Chlamydia infections in primary endometrial organoids. This foundation will provide a framework for future studies that target host and pathogens to better identify how Chlamydia subverts epithelial biology and the innate immune response.
Materials and Methods
Ethics statement
All animal experiments were approved and performed in accordance to the Duke University Institutional Animal Care and Use Committee.
Cell lines and conditioned medium
Vero cells were purchased from ATCC (CCL-81) and cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; Life Technologies) containing 10% fetal bovine serum (FBS; Sigma-Aldrich). L-WRN cells were purchased from ATCC (CRL-3276) and cultured in DMEM containing 0.5 mg/mL geneticin (Gibco) and 0.5 mg/mL hygromycin B (ThermoFisher) at 37°C with 5% CO2. Prior to generating conditioned medium, the cells were passaged twice in media without antibiotics. Conditioned medium was generated as previously described (Miyoshi and Stappenbeck, 2013). In brief, cells were plated in T175 tissue cultured treated flasks and grown to greater than 90% confluence. 25 mL of 1:1 DMEM/F12 (Gibco) media was collected every 24 hours, centrifuged at 500 x g for 5 min, and stored in 4°C. At the end of five days, the media were combined, sterile filtered, and stored in −80C.
Chlamydia strains, propagation, and transformation
C. trachomatis strains L2/434/Bu (CTL2; ATCC VR-902B), the genetically-modified M007, M407, and M923 chemical mutants (Kokes et al., 2015; Sixt et al., 2017) were propagated in Vero cells, harvested at 44-48 hpi by water lysis, sonication, diluted in SPG (sucrose-phosphate-glutamate) buffer to 1x concentration (75 g/l sucrose, 0.5 g/l KH4PO4, 1.2 g/l Na2HPO4, 0.72 g/l glutamic acid, pH 7.5), and stored as single use aliquots at −80°C. The C. muridarum strain CM006 was a gift from Catherine O’Connell (University of North Carolina), and was propagated in Vero cells and harvested at 36 hpi as above. The CM006 strain was transformed with pNigg-SW2-GFP as follows: Approximately 107 IFU were incubated with 10 μg DNA in buffer containing 0.9 mM calcium chloride for 30 min, added to confluent Vero cells in a six well plate, and centrifuged at 3,000 rpm for 30 min at 10°C. At 12 hours post-infection, 1 U/mL penicillin was added. The infections were passaged every 36 hours until inclusions were present and fluorescent. Transformants were subsequently plaque-purified to obtain a clonal strain.
To generate the spectinomycin-resistant TepP-deficient C. trachomatis, CTL2 was transformed using the TargeTron gene disruption system (Sigma-Aldrich) as the previously described TepP mutant (Carpenter et al., 2017) with the exception that an aadA cassette was inserted into the targeting site (between amino acids 821-822). Transformants were expanded in Vero cells in the presence of 150 ug/mL spectinomycin as described above, plaque purified, and verified by PCR analysis using primers flanking the insertion site and the aadA cassette. The ΔtepP::aadA strain was complemented with the E.coli-Chlamydia shuttle vector p2TK2-SW2 (Agaisse and Derré, 2013) containing a bla cassette and the TepP open reading frame containing its upstream promoter sequence. In brief, 108 IFU were incubated with 10 μg DNA in buffer containing 0.9 mM calcium chloride for 30 min, added to confluent Vero cells in a six well plate, and centrifuged at 3,000 rpm for 30 min at 10°C. At 12 hours post-infection, 1 U/mL penicillin was added. The infections were passaged every 48 hours until inclusions were present where penicillin concentration was increased to 10 U/mL. Transformants were subsequently plaque-purified in the presence of both spectinomycin and penicillin to obtain a clonal strain and verified by PCR using primers targeting both the aadA and bla cassette and the full-length tepP sequence.
Western Blots
For western blots, Vero cells were infected with the indicated strains for 48 hours, water lysed, sonicated, and boiled in Laemelli buffer for 10 min. The lysates were centrifuged at 9,600 x g for 5 min and equal volumes were resolved using a 10% SDS-PAGE gel. The resolved proteins were transferred to a nitrocellulose membrane (Amersham), blocked with a PBS solution containing 5% milk for one hour at 25°C, and sequentially probed with primary anti-TepP (1:500; (Chen et al., 2014)) and anti-Slc1 (1:1000; (Chen et al., 2014)) and the LiCOR infrared-conjugated secondary antibodies (1:10,000). Membranes were imaged using an Odyssey LiCOR.
EMO generation from the mouse endometrium and hormone stimulation
The C57/BL6J and B6.129(Cg)-Gt(ROSA)26Sor^tm4(ACTB-tdTomato,-EGFP)Luo/J (Strain no. 007676) mouse strains were purchased from Jackson Laboratories. The ZO-1 GFP knock-in mouse line was a generous gift from Dr. Terry Lechler (Duke University). The endometrium from ~ 6-8 week old females were dissected, washed in cold Dulbecco’s phosphate-buffered saline (PBS; Gibco) on ice, cleaned of vascular and fat tissue, minced into ~ 2 mm pieces, transferred to DMEM (Gibco) containing 0.2% collagenase A (Thermo), 10% FBS, and 1U/mL penicillin/streptomycin, (Gibco) then incubated on a shaker at 37°C for 2.5 – 3 hours. The tissue was washed three times with cold PBS and mechanically disrupted by shaking in cold 10 mL PBS containing 0.1% bovine serum albumin (BSA; Fraction V, Equitech-Bio) for 1 min. After allowing the tissue to settle for one minute, the supernatant was collected and passed through a 70 μm strainer (Falcon). The mechanical disruption was repeated once more, and the strainer was subsequently inverted over a new 50 mL conical tube and the epithelia were washed off the strainer three times with 10 mL PBS containing 0.1% BSA. The epithelial fraction was centrifuged at 500 x g for 5 min at 10°C, resuspended in cold DMEM/F12 (Gibco), mixed with low-hormone Matrigel (Corning) at a 1:1 ratio, and pipetted in 35 μL drops in a 24 well plate or 125 μL drops in a 35 mm glass-bottom dish (for microinjections). The plates were incubated at 37°C for 40 min before gently overlaying 2 mL 1:1 DMEM/F12 media containing 50% L-WRN conditioned media, 50 μg/mL gentamicin (Gibco), and 50 ng/mL EGF (StemCell Technologies). The media was changed every 2-3 days. For hormonal stimulation studies, EMOs were cultured in the presence of 17-β-estradiol (E2; Sigma-Aldrich) for four days.
Organoid microinjection
Microinjections were performed using an Eppendorf FemtoJet 4x coupled with a Stereo Microscope (Nikon). Chlamydia strains were diluted in PBS to a final concentration of 5E5-5E6 IFU, vortexed for 30s, and pipetted into a glass needle. Organoids were punctured once using a steep vertical angle and manually injected with equal volumes. When organoids were injected alone or co-injected with 3kD Texas-Red dextran (Invitrogen), dextran was used at a final concentration of 0.01 mg/mL.
Primary neutrophil isolation, co-culture with infected EMOs
To isolate primary neutrophils, mouse femurs were dissected, cleaned with an ethanol-soaked Kim wipe to remove other tissue, dipped in 70% ethanol then twice in RPMI media (Gibco). Both ends of the femur were cut and 10 mL RPMI media was passed through the bone with a 25G needle (BD Biosciences). The supernatant was centrifuge at 600 x g for 5 minutes, resuspended in 5 mL red cell lysis buffer (Millipore), and incubated for 5 minutes. An additional 5 mL of RPMI media was added to the cells and centrifuged at 600 x g for 5 minutes. The cells were resuspended gently in 0.5 mL RPMI media before following the negative selection EasySep™ Mouse Neutrophil Enrichment Kit (StemCell Technologies). Neutrophils were labeled with 5 μM CellTracker (CFMDA; ThermoFisher) for 10 minutes at 37°C, centrifuged at 300 x g for 10 minutes at 4°C, and resuspended in 0.5 mL cold RPMI. Neutrophils were counted and 3E5 cells were added directly to the media and cultured for an additional 20 hours.
Antibodies
For immunofluorescence microscopy, the following antibodies were used: mouse anti-GM130 (BD Biosciences, Cat no. 610822; 1:200), anti-alpha-tubulin (Sigma, Cat no. T5618, Clone B-5-1-2; 1:200), anti-Muc1 (Cell Signaling Technologies, Cat no. VU4H5; 1:100), anti-β-catenin (BD Biosciences, Cat no. 610153 Clone 14; 1:400), anti-MOMP (Santa Cruz, Cat no. 57678; 1:500), anti-pan-Keratin (Sigma-Aldrich Cat no. C2562; 1:100), and rabbit anti-E-cadherin (Cell Signaling Technologies; Cat no. 3195S, Clone 24E10; 1:200), anti-acetylated-alpha-tubulin (Cell Signaling Technologies, Cat no. 5335S 1:200), anti-SEPT2 (Protein Tech; Cat no. 11397-1-AP; 1:100), anti-Cap1 (Gift from A. Subtil; 1:250).
Brightfield and immunofluorescence microscopy
Organoid growth was monitored by brightfield microscopy using the EVOS FL Cell Imaging System (ThermoFisher) equipped with 2x/0.06 and 10x/0.25 NA objectives and a CCD camera. For fixed samples, organoids were rinsed twice with warm PBS and incubated with warm PBS containing 3% formaldehyde (Sigma) for 20 minutes. The fixative was removed gently, and the organoids were resuspended in 0.25% ammonium chloride (Sigma-Aldrich) and transferred to a 1.5 mL tube, centrifuged at 500 x g for 5 min, resuspended gently in 2% BSA (Sigma-Aldrich) containing 0.1% triton x-100 and incubated with gentle rocking for 30 minutes. Organoids were centrifuged again at 500 x g for 5 min, incubated with the indicated primary antibodies diluted in 0.5 mL 2% BSA containing 0.1% triton x-100, and incubated at 25°C for 2-3 hours or overnight at 4°C with gentle rocking. Organoids were washed once with 2% BSA, centrifuged at 500 x g for 5 min, and incubated with secondary antibodies diluted in 1.0 mL 2% BSA containing 0.1% triton x-100 for 1.5 hours at 25°C with gentle rocking. Acti-stain conjugated to Alexa-555 (1:500; Cytoskeleton, Inc) and Hoechst (2 μg/mL; ThermoScientific) were added for the final 20 minutes of incubation with the secondary antibodies. The organoids were centrifuged at 500 x g for 5 min and resuspended in 30 μL Vectashield (Vector Labs; H-1000) using a cut pipet tip and pipetted onto a coverslide. A coverslip was overlaid gently and sealed with nail polish.
Organoids were imaged using an inverted confocal laser scanning microscope (LSM 880; Zeiss) equipped with a motorized stage, Airyscan detector (Hamamatsu), and diode (405 nm), argon ion (488 nm), double solid-state (561 nm), and helium-neon (633 nm) lasers. Images were acquired using a 20x/0.8 NA air or 40x/1.2 NA water objective (Zeiss) and deconvolved using automatic Airyscan Processing in the Zen Software (Zeiss). Images were opened in ImageJ (NIH) or the Imaris software and exported TIFFs were rendered in the Adobe suites (Photoshop and Illustrator). Only linear adjustments were made to fluorescence intensity.
Time-lapse microscopy of inclusion dynamics and neutrophil recruitment
Organoids cultured in 35 mm glass bottom dishes (Mat-Tek) were imaged live using an inverted microscope (Zeiss AxioObserverZ.1) equipped with a motorized stage containing a heated Insert P environmental chamber (Zeiss), XLIGHT V2 spinning disk unit (Crest Optics), and an ORCA Flash 4.0 V3 camera (Hamamatsu). Images were acquired using a 20x/0.8 NA objective (Olympus), an LDI multiline laser (89 North) using two micron sections at 6-10 minute intervals for 16-18 hours. Using the same microscope, neutrophil recruitment to infected EMOs was imaged using five micron confocal sections and spanning ~ 100 μm above and below each organoid. Timelapse microscopy of neutrophil motility was performed using a 20x/0.8 NA objective (Olympus) or a 60x/1.4 NA objective (Olympus). Confocal sections (2 μm) were acquired every three minutes for the duration of three hours. All images were rendered in ImageJ (NIH). Exported TIFFs were reconstructed in the Adobe suites (Photoshop, Illustrator).
Image analysis
To quantify EMO size, microscopy images were imported into ImageJ and converted to 8-bit TIFFs. EMO borders were identified using the find edges algorithm and converted to a binary image. Using the find maxima algorithm, each EMO was identified, the area exported into Excel and plotted in R.
Golgi reorganization around the inclusion was quantified in ImageJ. Microscopy images were imported and converted to 8-bit TIFFs. The inclusion edges were manually traced to measure the perimeter length. Using the segmented line tool, the Golgi signal at the inclusion was also traced to measure its length. Each value was imported into Excel to generate the percent distribution around the inclusion perimeter and subsequently plotted in R.
To quantify cell death, microscopy images of EMOs incubated with propidium iodide were quantified in ImageJ. Microscopy images were imported and converted to 8-bit TIFFs. Maximum projection images were background subtracted using the rolling ball radius (value = 50) and puncta were identified using the find maxima algorithm.
Neutrophil recruitment to infected EMOs was quantified in ImageJ. Maximum projection images were background subtracted as above. To identify proximal neutrophils, a circle was placed around the inclusion 200 μm from the EMO basolateral edge. The number of neutrophils were identified using the find maxima algorithm, imported into Excel and plotted in R.
Competing interests
R.H. Valdivia is co-founder at Bloom Sciences (San Diego, CA). The company did not sponsor any of the shown work nor has financial interests in the outcomes of these studies.
Funding
This work was supported by National Institutes of Health (F32AI138371 to L.D.) and (AI-R01134891 to R.H.V).
Acknowledgements
We thank Dr. Lisa Cameron at the Duke Light Microscopy Core Facility and Dr. David Tobin for assistance with microscopy, and members of the Valdivia lab for critical feedback on this project. Dr. Jorn Coers and Dr. Ryan Finethy for assistance with mice and immune cell isolations, Dr. Terry Lechler for the ZO-1 GFP knock-in mice, and Dr. Agathe Subtil for the Cap1 antibody.