The host adherens junction molecule nectin-1 is downregulated in Chlamydia trachomatis-infected genital epithelial cells

The obligate intracellular bacterium Chlamydia trachomatis is a significant Gram-negative human pathogen, with four million new cases per year in the USA. Long-term C. trachomatis genital tract infections are often asymptomatic and associated with severe ascending complications, including epididymitis, prostatitis, ectopic pregnancy, salpingitis and infertility (Darville, 2000). The chlamydial developmental cycle begins when the elementary body (EB) binds to and enters mucosal epithelial cells. Once internalized, the EB remains within an inclusion and differentiates into a reticulate body (RB). Using host cell nutrients, the RB grows, divides by binary fission and eventually produces more EBs, which are released by host-cell lysis (Wyrick, 2000). When developing chlamydiae are confronted with nutrient deficiency or penicillin exposure, they deviate from the normal developmental cycle and enter into a state called persistence, which is characterized by formation of an aberrantly enlarged, viable but non-infectious chlamydial RB (Hogan et al., 2004). Notably, the chlamydiae can re-enter and complete the normal developmental cycle once the persistence inducer is removed.

Upon infection, chlamydiae alter host cellular functions in a variety of ways. Chlamydial infection prevents host cell apoptosis, induces reorganization of the actin cytoskeleton and alters host cellular signalling mechanisms (Hackstadt, 1999; Hatch, 1999). C. trachomatis infection also disturbs cell–cell contacts between polarized epithelial cells in culture (Wyrick et al., 1989). This cell–cell disassociation was later shown to be due, at least in part, to disruption of N-cadherin-dependent cell–cell junctions and breakdown of the N-cadherin/β-catenin complex (Prozialeck et al., 2002). Notably, infected epithelial cell release has also been observed in vivo (Doughri et al., 1972; Soloff et al., 1985). Doughri et al. (1972) further suggested that release of chlamydiae within intact host cells might protect the organisms from anti-chlamydial antibodies. In both studies, the authors hypothesize that polymorphonuclear neutrophils (PMNs) responding to chlamydial infection promote detachment of chlamydia-infected cells from the mucosal surface, resulting in the release of intact, infected cells into the cervical lumen (Doughri et al., 1972; Soloff et al., 1985). The seminal observation that chlamydial infection in the absence of PMNs loosens cell–cell junctions by promoting breakdown of the N-cadherin/β-catenin complex (Prozialeck et al., 2002) suggests that the developing chlamydiae may also contribute directly to the release observed in vivo of infected cells from the infected mucosa.

In epithelial cells, junctional complexes, which are composed of tight junctions (TJs), adherens junctions (AJs) and desmosomes (DSs), play critical roles in forming intercellular adhesions. Intercellular adhesions, in turn, are intimately involved in maintaining cell structure, transmitting information, regulating cellular migration, and imparting strength and rigidity to tissues (Farquhar & Palade, 1963; Gumbiner, 1996; Lauffenburger & Horwitz, 1996; Takeichi, 1995). Importantly, the organization and function of TJs and DSs are dependent on the formation and maintenance of AJs (Tsukita & Furuse, 1999). AJs are located at the basal side of TJs and connect the lateral surfaces of adjacent epithelial cells. AJs form complexes with actin and myosin filaments to internally brace cells and thereby control their shape. E-cadherin and claudin are two well-studied cell–cell adhesion molecules found in AJs and TJs, respectively.

A novel cell–cell adhesion system consisting of nectin and afadin has recently been identified at AJs in epithelial cells, neurons and fibroblasts (Takai & Nakanishi, 2003). Nectins are Ca²⁺-independent, Ig-like cell–cell adhesion molecules with three extracellular Ig-like loops, a single transmembrane region (except for nectin-1γ) and a cytoplasmic tail. The nectin family currently contains four members (nectins 1–4) (Sakisaka & Takai, 2004; Takai & Nakanishi, 2003). Afadin is a nectin- and F-actin-binding protein that connects the transmembrane domain of nectin to the actin cytoskeleton (Takai & Nakanishi, 2003). This nectin-based adhesion system is required not only for the formation of AJs but also for the subsequent formation of TJs (Takai & Nakanishi, 2003). Furthermore, activation of Cdc42 and Rac small G proteins through c-Src by trans-interaction of nectins has been implicated in controlling cell–cell adhesion and cell polarization (Takai et al., 2003).

In this study, Western blot analyses demonstrate that accumulation of host nectin-1 is significantly decreased in C. trachomatis serovar E-infected cervical epithelial cells. This downregulation requires active C. trachomatis protein synthesis and/or replication. RT-PCR analyses further show that reduced expression of nectin-1 likely occurs post-transcriptionally. Because the nectin–afadin adhesion system is critical for formation of functional E-cadherin-based AJs and subsequent claudin-based TJs, and because chlamydial infection has been shown previously to disrupt cell–cell junctions (Prozialeck et al., 2002), our data suggest that C. trachomatis may disrupt AJs, at least in part, by diminishing nectin-1 expression.

Cell culture, chlamydial infection and titrations.
HeLa cells, a human cervical adenocarcinoma epithelial cell line [American Type Culture Collection (ATCC) no. CCL2], were cultured as described previously (Deka et al., 2006). A human urogenital isolate of C. trachomatis E/UW-5/CX was originally obtained from S. P. Wang and C. C. Kuo (University of Washington, Seattle, WA). The same standardized inoculum of C. trachomatis serovar E EBs, propagated in McCoy cells, was used for all experiments (Wyrick et al., 1996). For infection experiments, HeLa cells either were grown on standard tissue culture plates or were polarized on 3.0 µm, collagen IV-coated chamber inserts (Biocoat 4544, Becton Dickinson), as described elsewhere (Wyrick et al., 1989). Host cells were infected at an m.o.i. of 1 using crude EB stock, which infects ∼90 % of the HeLa cells, for 1 h at 35 °C. In some experiments, HeLa cells were infected with an equivalent amount of UV-inactivated C. trachomatis serovar E (C. trachomatis-UV). Mock-infected host cells were treated similarly, except that they were exposed to 200 µl 2SPG (0.2 M sucrose, 6 mM NaH₂PO₄, 15 mM Na₂HPO₄, 5 mM L-glutamine, pH 7.2). After infection, the inoculum was aspirated and the infected cells were re-fed with growth medium and incubated at 35 °C for the indicated times. Chlamydial titrations were performed as described previously (Deka et al., 2007). Each experiment was repeated three times and each repeat contained three biological replicates.

Generation of C. trachomatis-UV.
A UV cross-linker (Spectrolinker XL-1500, Spectronics Corporation) was used to generate stocks of UV-inactivated, replication-incompetent C. trachomatis (C. trachomatis-UV). Stock C. trachomatis serovar E was thawed on ice and 200 µl was aliquoted into each well of a 24-well plate. The plate was placed on a 4 °C heat sink during UV exposure to prevent heat inactivation of the samples. EB inactivation was assayed by performing chlamydial titrations (Deka et al., 2007). A UV dose of 1.0 J cm^–2 was sufficient to completely inactivate undiluted C. trachomatis stocks (data not shown).

RNA and DNA isolation.
Total RNA and DNA were isolated from mock- or C. trachomatis-infected HeLa cells at 48 and 60 hours post infection (h.p.i.) using the method described previously by Deka et al. (2006). Total RNA and DNA preparations were quantified by measuring OD at 260 and 280 nm. All samples had OD₂₆₀/OD₂₈₀ ratios >1.9. The quality and concentration of each RNA sample were further confirmed by analysis on a 2100 Bioanalyser instrument (Agilent) using the RNA 6000 Nano LabChip kit. All samples had RNA integrity numbers (RINs) between 9.0 and 10.0 (data not shown).

PCR, reverse transcription and RT-PCR.
Total cellular DNA was used as a template to amplify human glyceraldehyde-3-phosphate dehydrogenase (GAPDH) for determination of host genome copy number. Total cellular RNA samples were subjected to reverse transcription as described previously (Deka et al., 2006), except that 2 µg total RNA was used. RT-PCR reactions were then carried out using RT(+) and RT(–) cDNA reactions as templates, as appropriate. Experimental template DNAs/cDNAs were used in PCR reactions at dilutions ranging from 1/10 to 1/1000 [in double-distilled H₂O (ddH₂O)], such that each reaction was in the linear amplification range. The amplification conditions were based on those described previously (Deka et al., 2006). Published primer sets included those for human β-actin (Nelson et al., 2005) and human GAPDH (Deka et al., 2006). Specific primers and positive oligonucleotide amplification controls for human nectin-1α, nectin-1β and nectin-1γ were designed using Oligo 4.0 software. These are listed in Supplementary Table S1. Most reactions were performed using the following cycling conditions: 94 °C, 1 min; 60 °C, 1 min; 72 °C, 1 min for 35 cycles. Human β-actin reactions were performed under identical conditions but for 30 cycles; human GAPDH PCR reactions were performed for 35 cycles and annealed at 67 °C. The resulting PCR products were separated by electrophoresis on 2.5 % agarose/Tris-buffered EDTA (TBE) gels and stained with ethidium bromide. A Bio-Rad Chemi Doc XRS image capture system with Quantity One V4.5.0 software was used to visualize and quantify amplimers.

SDS-PAGE and Western blotting.
Monolayers were lysed and denatured as described previously (Deka et al., 2006). Samples were then normalized for total protein content using SYPRO Ruby stain (Bio-Rad) according to the manufacturer's instructions. Samples were diluted to identical concentrations, subjected to SDS-PAGE and Western blotted (Deka et al., 2006). Primary antibodies used included mouse monoclonal anti-nectin-1 CK6 (1 : 200 dilution; sc-21722, Santa Cruz Biotechnology), mouse monoclonal anti-nectin-1 CK8 (1 : 200 dilution; 37-5900, Invitrogen–Zymed), rabbit polyclonal anti-human focal adhesion kinase (FAK) C-20 (1 : 2000 dilution; sc-558, Santa Cruz Biotechnology) and goat polyclonal anti-chlamydial major outer-membrane protein (MOMP; 1 : 5000 dilution; B65266G, BioDesign International). Bound primary antibodies were detected using the corresponding secondary antibodies conjugated to horseradish peroxidase and visualized using SuperSignal West Pico reagent (Pierce). Specific bands were quantified using an FX phosphorimager and Quantity One V2.5.0 software (Bio-Rad). To control for small variations in cell number and gel loading between sample lanes, the quantity of nectin-1 in each sample was normalized to the amount of FAK protein detected in that same lane. Previous studies from our laboratory have demonstrated that FAK accumulation is not altered by chlamydial infection and therefore makes an ideal internal control (Deka et al., 2006).

Statistical analyses.
Statistical analyses were performed using Microsoft Excel. A two-sample t test for independent samples was used for comparison of means. Significance was defined as P≤0.05.

Nectin-1 accumulation is reduced in C. trachomatis-infected HeLa cells
Nectin-1 is an important cell–cell adhesion molecule involved in the formation and maintenance of AJs and TJs. Previous studies have demonstrated that cell–cell contacts are disrupted in C. trachomatis-infected cervical epithelial cells (Prozialeck et al., 2002). To determine whether chlamydial infection modulates nectin-1 expression, monolayers of non-polarized HeLa cells were either mock- or C. trachomatis serovar E-infected at an m.o.i. of 1. Cell lysates were collected 48 h.p.i. and processed for SDS-PAGE analyses. Duplicate gels were either stained with SYPRO Ruby (data not shown) or Western blotted (Fig. 1a) using anti-nectin-1 CK6 mAb, anti-C. trachomatis MOMP or anti-FAK, as described previously (Deka et al., 2006). Nectin-1 bands were quantified, normalized to FAK and plotted in Fig. 1(b). FAK was chosen as an internal control protein because its accumulation is not altered by chlamydial infection (Deka et al., 2006). As expected, MOMP expression was detected in C. trachomatis-infected cells alone. Nectin-1 accumulation was decreased by up to 85 % in C. trachomatis-infected HeLa cells compared with that in mock-infected controls. Sequential immunoprecipitation and blotting experiments with a different nectin-1-specific mAb (Supplementary Fig. S1b) confirmed that the protein detected on the blots in Fig. 1 was indeed nectin-1. Additionally, little or no nectin-1 was immunoprecipitated by either antibody from chlamydia-infected cells harvested at 48 h.p.i., confirming the observations described above.

(18K): Fig. 1. Nectin-1 accumulation is reduced in C. trachomatis-infected HeLa cells. HeLa cells were either mock infected (Mock inf.) or C. trachomatis serovar E infected (Ct inf.) at an m.o.i. of 1. Cell lysates were collected 48 h.p.i. and Western blotted using anti-nectin-1 CK6, anti-C. trachomatis MOMP or anti-FAK (C-20). Nectin-1 bands were quantified, normalized to the internal control protein FAK and plotted in (b). Results are expressed as the mean±SEM of three biological replicates. Significant (P≤0.05) difference from the mock-infected controls is indicated by an asterisk. The data are representative of three independent experiments.

In vivo, polarized epithelial cells differentially distribute proteins and lipids to the plasma membrane, creating two distinct surfaces, the apical and basolateral domains. Previous studies have shown that polarized and non-polarized cells differ in C. trachomatis infection efficiency, duration of the C. trachomatis developmental cycle, and their response to IFN-γ (Arno et al., 1990; Kane & Byrne, 1998; Tam et al., 1992). Thus, we wanted to investigate whether or not the polarity of epithelial cells would affect the observed nectin-1 reduction. Western blotting analyses (Supplementary Fig. S2a) showed that nectin-1 was also downregulated in C. trachomatis-infected polarized cells. At 48 h.p.i., nectin-1 accumulation was decreased by as much as 88 % in C. trachomatis-infected polarized HeLa cells compared with that in mock-infected controls (Supplementary Fig. S2b). Because similar results were obtained from both non-polarized and polarized HeLa cells, non-polarized cells were used in subsequent experiments.

Nectin-1 accumulation is reduced in C. trachomatis-infected HeLa cells from 36 to 60 h.p.i
To more precisely define when downregulation of nectin-1 occurs during the C. trachomatis developmental cycle, an infection time-course ranging from 0 to 60 h.p.i. was conducted. HeLa cells were either mock- or C. trachomatis-infected. Cell lysates were collected at 0, 24, 36, 48 and 60 h.p.i. and Western blotted with (i) anti-nectin-1 (CK6), (ii) anti-FAK (C-20) and (iii) anti-MOMP. A representative blot is shown in Fig. 2(a); the quantification of this blot is shown in Fig. 2(b). The nectin-1 quantity observed in C. trachomatis-infected cells from three independent experiments was expressed as a percentage of that observed in mock-infected control cells at each time interval and plotted (Fig. 2c). These data demonstrate that C. trachomatis infection significantly reduced accumulation of nectin-1, starting at 36 h.p.i., and that the reduction was maximal at 60 h.p.i.

(35K): Fig. 2. Nectin-1 accumulation is reduced in C. trachomatis-infected HeLa cells from 36 to 60 h.p.i. (a) Cell lysates from mock-infected (Mock inf.) or C. trachomatis-infected (Ct inf.) HeLa cells were collected at 0, 24, 36, 48 and 60 h.p.i. and Western blotted using anti-nectin-1 CK6, anti-C. trachomatis MOMP or anti-FAK (C-20). (b) The quantity of nectin-1 detected in panel (a) was normalized to the internal control protein FAK and plotted. (c) The quantity of Nectin-1 observed in C. trachomatis-infected cells is expressed as a percentage of that observed in mock-infected control cells at each time interval. Results are expressed as the mean±SEM of three biological replicates. Groups significantly (P≤0.05) different from C. trachomatis-infected cells at 0 h.p.i. are indicated by asterisks. The data are representative of three independent experiments.

Nectin-1 downregulation requires C. trachomatis protein synthesis and/or replication
To extend these findings, we next investigated whether C. trachomatis protein synthesis and/or replication was required for nectin-1 downregulation. Nectin-1 expression in persistently infected cells was also examined. HeLa cells were either mock- or C. trachomatis-infected and re-fed immediately after infection with growth medium plus ddH₂O, medium plus 60 µg chloramphenicol ml^–1, which specifically inhibits the function of bacterial 50S ribosomal subunits, or medium plus 20 U penicillin G ml^–1, a known inducer of chlamydial persistence. Cell lysates were harvested at 48 h.p.i. for Western blot analyses (Fig. 3a, b). The quantity of nectin-1 observed in C. trachomatis-infected cells from three independent experiments was expressed as a percentage of that observed in mock-infected control cells under the same experimental conditions (Fig. 3c). As a control to confirm that each antibiotic was working as expected, duplicate cultures from each drug-exposure group were subjected to chlamydial subpassage titration analyses. The numbers of inclusion-forming units (IFU) in the undiluted inocula were then calculated and expressed as IFU ml^–1 (Fig. 3d). Nectin-1 downregulation in C. trachomatis-infected cells was prevented by chloramphenicol exposure (Fig. 3a, lanes 5 and 6), demonstrating that C. trachomatis protein synthesis and/or replication is required for this effect. Nectin-1 levels also decreased in the presence of penicillin G (Fig. 3a, lanes 7 and 8), although to a lesser degree than that observed in diluent-exposed, chlamydiae-infected monolayers (Fig. 3b, c). These data indicate that nectin-1 accumulation can also be reduced by penicillin-induced, persistent C. trachomatis infection. Chlamydial subpassage experiments demonstrated that both chloramphenicol and penicillin G exposure significantly reduced recovery of infectious EBs, as expected (Fig. 3d).

(37K): Fig. 3. C. trachomatis protein synthesis and/or replication is/are required for nectin-1 downregulation. (a) Cells were re-fed immediately after C. trachomatis infection with growth medium+ddH2O, 60 µg chloramphenicol ml–1 (camp) or 20 units penicillin G ml–1 (penG). Cell lysates were harvested at 48 h.p.i. and Western blotted using anti-nectin-1 CK6, anti-C. trachomatis MOMP or anti-FAK (C-20). (b) The quantities of nectin-1 detected on the blot in panel (a) were normalized to FAK and plotted. (c) The quantity of nectin-1 observed in C. trachomatis-infected cells is expressed as a percentage of that observed in mock-infected control cells under that set of experimental conditions. The mean±SEM from three biological replicates is shown. Groups significantly (P≤0.05) different from chloramphenicol-exposed, C. trachomatis-infected cells are indicated by asterisks. (d) Duplicate cell cultures from each drug-exposure group were subjected to chlamydial subpassage titration. Results are expressed as the mean±SEM of three biological replicates. Groups significantly (P≤0.05) different from non-drug-exposed, C. trachomatis-infected cells are indicated by asterisks. All data shown here are representative of three independent experiments.

Viable C. trachomatis EBs are required for nectin-1 downregulation
To determine whether viable C. trachomatis EBs are required for downregulation of nectin-1, UV-inactivated C. trachomatis was used to infect HeLa cells. Previous studies have demonstrated that UV-inactivation of C. trachomatis renders the bacteria completely replication-incompetent (Eissenberg et al., 1983). HeLa cultures were mock-, C. trachomatis- or C. trachomatis-UV-infected, collected at 48 h.p.i. and processed for Western blot analyses (Fig. 4a). Average nectin-1 accumulation was quantified, normalized and plotted (Fig. 4b). Nectin-1 accumulation was reduced in C. trachomatis-infected cells but not in cultures infected with an equivalent quantity of UV-inactivated C. trachomatis EB, demonstrating that nectin-1 is downregulated only if cells are infected with viable EBs.

(21K): Fig. 4. Viable C. trachomatis EBs are required for nectin-1 downregulation. (a) HeLa cells were mock (Mock), C. trachomatis (Ct) or C. trachomatis-UV (Ct-UV) infected. Cells were collected at 48 h.p.i. and proteins were Western blotted using anti-nectin-1 CK6, anti-C. trachomatis MOMP or anti-FAK (C-20). (b) The quantity of nectin-1 was normalized to the internal control protein FAK and plotted. Results are expressed as the mean±SEM of three biological replicates. Groups significantly (P≤0.05) different from the mock-infected controls are indicated by asterisks. The data shown are representative of three independent experiments.

C. trachomatis infection of HeLa cells does not alter accumulation of nectin-1α, β and γ transcripts
Nectin-1 has three splice variants: nectin-1α, nectin-1β and nectin-1γ. All three nectin-1 isoforms encoded by these transcripts associate with other adhesion molecules and play important roles in the formation of cell–cell junctions. Although the nectin-1α and nectin-1β proteins both contain the CK6 and CK8-reactive epitopes (Supplementary Fig. S1a), the observed molecular mass (75 kDa) is most consistent with the reported size for nectin-1α (Struyf et al., 2005). To aid in elucidating the mechanism by which host cell nectin-1 expression is altered in response to C. trachomatis infection, we investigated whether downregulation of nectin-1 occurs at the transcriptional level. Total RNA from mock- or C. trachomatis-infected cells was isolated and subjected to RT-PCR using primers specific for human β-actin, nectin-1α, nectin-1β and nectin-1γ transcripts. Parallel total DNA samples were amplified using GAPDH-specific primers to determine host genome copy number. All amplimers were the expected size (Fig. 5a) and the identity of each was confirmed by DNA sequencing (data not shown). In each experiment, a six-log dilution series of known DNA template controls was used to generate standard curves for amplification. Experimental samples were only quantified if they fell within the linear range of the PCR. Amplification products were not observed in template-negative samples (Fig. 5a, lane 1) or in RT(–) controls (data not shown). The quantity of nectin-1α, β and γ products was similar in mock- compared with C. trachomatis-infected samples at both 48 and 60 h.p.i. (Fig. 5a, lanes 2–5). All amplimers were quantified, normalized to host genome copy number, as ascertained by PCR with human GAPDH-specific primers, and plotted in Fig. 5(b). Statistical analyses showed that there was no significant difference in accumulation of any of the nectin-1 transcripts in mock- versus C. trachomatis-infected cells (Fig. 5b). These data indicate that expression of nectin-1 is not regulated at the transcriptional level and thus is probably regulated post-transcriptionally.

(42K): Fig. 5. C. trachomatis infection of HeLa cells does not alter accumulation of nectin-1α, β and γ transcripts. (a) HeLa cells were mock (Mock) or C. trachomatis (Ct) infected at an m.o.i. of 1. Total RNA was isolated at 48 or 60 h.p.i., as described in Methods, and subjected to RT-PCR analyses using transcript-specific oligonucleotide probes. Nectin-1α (117 bp), nectin-1β (105 bp), nectin-1γ (75 bp) and β-actin (230 bp) amplimers were electrophoresed and photographed. Representative photographs are shown. The positions of DNA size markers are shown to the left of each gel image in bp. Neg, control PCR containing no DNA template. (b) Specific DNA amplimers were quantified as described in Methods. Nectin-1α (black bars), nectin-1β (grey bars), nectin-1γ (white bars) and β-actin (diagonally striped bars) amplimer quantities were normalized to host genome copy number, as ascertained by PCR with human GAPDH-specific primers (data not shown). The values plotted are means±SEM of three independent experiments. The value obtained for each independent experiment is the mean obtained from three biological replicates.

Chlamydiae have evolved various strategies to avoid host defence systems and complete their intracellular growth cycle. They have acquired specific mechanisms to prevent productively and persistently infected cells from undergoing apoptosis (Dean & Powers, 2001; Dong et al., 2005; Fan et al., 1998; Fischer et al., 2004; Geng et al., 2000; Pirbhai et al., 2006; Rajalingam et al., 2001), which may allow chlamydiae to maintain long-term survival within infected cells. Chlamydiae also alter host cell–cell contacts and signal transduction (Prozialeck et al., 2002; Xia et al., 2003), which may facilitate chlamydial growth. Finally, chlamydiae also modify the host cell cytoskeleton (Dong et al., 2004; Majeed et al., 1993), which may benefit chlamydial intravacuolar replication. In the present study, we observed a remarkable loss of nectin-1 in C. trachomatis-infected HeLa cells. We further demonstrated that active C. trachomatis protein synthesis and/or replication and viable C. trachomatis EBs are required for this effect. Lastly, we also noted that persistent chlamydial infection, induced by penicillin G, also downregulates nectin-1 accumulation. Because different persistence inducers act upon the developing chlamydiae by divergent mechanisms, we must examine other stressors to determine whether the nectin-1 decrease is a general characteristic of persistently infected epithelial cells.

It is currently accepted that nectin-1 plays a critical role in the formation and maintenance of AJs and TJs. Disruption of nectin-1 would inevitably affect the integrity of cell–cell junctions, which may be implicated in chlamydial pathophysiological effects on the host. This notion is consistent with a recent study showing that cervical epithelial cells separate from each other as a consequence of C. trachomatis infection (Prozialeck et al., 2002). Interestingly, the important human pathogens Helicobacter pylori, Shigella flexneri and Salmonella typhimurium also downregulate essential components of adhesion and tight junctions, including E-cadherin, claudin-1 and ZO-1. This leads to decreased transepithelial electrical resistance and increased paracellular permeability, which facilitates cell-to-cell spreading of these pathogens (Jepson et al., 1995; Sakaguchi et al., 2002; Sears, 2000; Terres et al., 1998). Collectively, these findings suggest that disruption of adhesion and tight junctions to compromise cell–cell barriers may be an important strategy exploited by intracellular pathogens to manipulate host cells. Although C. trachomatis serovar E is not an invasive pathogen, it may diminish nectin-1 expression to aid lateral cell–to-cell spread. Surprisingly, in contrast to C. trachomatis and the pathogens mentioned above, Chlamydia pneumoniae upregulates the expression of adherens junction proteins VE-cadherin, N-cadherin and β-catenin and transiently downregulates the expression of the TJ protein occludin to alter junctional complexes, facilitating its transmission in human brain microvascular endothelial cells (MacIntyre et al., 2002). Why C. trachomatis and C. pneumoniae might use two distinct mechanisms to interact with junctional complexes is currently unknown.

While nectin-1 protein expression was decreased in C. trachomatis-infected HeLa cells, accumulation of nectin-1α, β and γ transcripts was unchanged. These data indicate that reduction of nectin-1 was not regulated transcriptionally, and thus is likely to be downregulated at the post-transcriptional level. Recently, several host proteins have been reported to undergo degradation upon C. trachomatis infection (Balsara et al., 2006; Dong et al., 2004, 2005; Fischer et al., 2004; Pirbhai et al., 2006; Ying et al., 2005; Zhong et al., 1999, 2000). The chlamydia-secreted protease CPAF (chlamydial protease/proteasome-like activity factor) is responsible for the degradation of many of these host proteins. Given the broad range of cellular proteins cleaved by CPAF, it is possible that CPAF also degrades nectin-1. Notably, there is little or no difference in CPAF expression between productive and persistent infections (Heuer et al., 2003; Shaw et al., 2002). Thus, the observation that nectin-1 was decreased in both C. trachomatis productively and persistently infected samples is consistent with the notion that CPAF degrades nectin-1. Alternatively, it is also possible that chlamydial infection induces a host-cell-derived protease or proteosomal activity that degrades nectin-1. The role of CPAF and other chlamydial or host-derived proteases in regulating nectin-1 accumulation in chlamydiae-infected cells requires further investigation.

Although the mechanism by which C. trachomatis downregulates nectin-1 expression is of significant interest, a more important question remains. Why does C. trachomatis reduce host cell nectin-1 accumulation? There are at least two plausible hypotheses. First, nectin-1 downregulation, and subsequent disruption of cell–cell adhesions, may facilitate host structural rearrangements required for inclusion enlargement or EB release at the end of the developmental cycle. This prediction is supported by the observation that nectin-1 accumulation starts to decrease at 36 h.p.i., which is mid-developmental cycle for C. trachomatis serovar E. The second, and perhaps more intriguing possibility, is that nectin-1 downregulation facilitates C. trachomatis dissemination within the host genital tract (Supplementary Fig. S3). In this case, weakening of contacts between an infected genital epithelial cell and adjacent, uninfected cells would allow release of the intact, infected cell into the genital tract lumen. The infected cell would then drift away from the original infection site in the genital mucus, aiding chlamydial dissemination in at least two ways. First, chlamydial EB within the exfoliated cell would be shielded from neutralization by secretory IgA, cationic anti-microbial peptides and other anti-bacterial compounds, as long as the host cell remained intact. Second, if the infected cell ruptured in close proximity to the genital mucosa, the local concentration of infectious EB would be very high, increasing the probability that mucosal epithelia in the vicinity would be successfully infected. In contrast, EB released from host cells still within the genital epithelial layer (Supplementary Fig. S3, right) would be both quickly diluted and immediately subject to neutralization by anti-bacterial compounds. Therefore, only host cells in close proximity to the release site would have a high probability of becoming infected. Thus, we envision that the released, inclusion-containing epithelial cell would act like a chlamydial cluster bomb, allowing delivery of a concentrated inoculum over a relatively long distance and increasing the possibility of a hit if the host cell ruptured near the genital epithelial cell layer. Interestingly, release of intact, chlamydiae-infected host cells from polarized monolayers in culture (Wyrick et al., 1989) as well as from infected epithelium in vivo has been observed (Doughri et al., 1972; Soloff et al., 1985). Of course, it is always possible that the observed nectin-1 decrease is an indirect side-effect of other host cytoskeletal or physiological alterations induced during chlamydial infection. Future studies will be directed toward determining which of these hypotheses is correct.

Finally, it should be noted that nectin also plays a critical role in intracellular signalling. Recent studies have demonstrated that Cdc42 and Rac small G proteins are activated by trans interactions of nectin in a PI3 kinase-independent manner in epithelial cells (Honda et al., 2003; Kawakatsu et al., 2002). Although the precise mechanisms of this activation remain to be elucidated, Cdc42 and Rac activities are required for the formation and maintenance of AJs in epithelial cells (Braga et al., 1997, 1999; Etienne-Manneville & Hall, 2002; Takaishi et al., 1997; Van Aelst & Symons, 2002). It is reasonable to speculate that the disruption of cell–cell junctions observed by Prozialeck et al. (2002) upon C. trachomatis infection results from insufficient activation of Cdc42 and Rac in response to reduced levels of nectin-1. Furthermore, activated Cdc42 and Rac stimulate the c-Jun N-terminal kinase (JNK) signalling pathway (Honda et al., 2003), which has been implicated in regulating cell growth, transformation and apoptosis (Johnson & Lapadat, 2002; Lin, 2003). As previously discussed, chlamydial infection strongly inhibits host cell apoptosis, and it is possible that reduced nectin-1/JNK signalling may play a role in this inhibition. Given the known interplay between nectin-1 and host cellular signalling pathways, dissecting the effect of C. trachomatis infection on nectin-mediated signal transduction events will be a key aspect of future experiments.

The authors would like to thank Dr Priscilla B. Wyrick and Dr Dennis M. Defoe for helpful discussion of these experiments and reviewing the manuscript. The authors also thank Dr John Laffan, Dr Sophie Dessus-Babus and Jennifer Vanover for their assistance on this project. This work was supported by NIH Public Health Service grant R21 AI59563 to R. V. S., East Tennessee State University (ETSU) research development committee (RDC) grant 04-024M to R. V. S., the James H. Quillen College of Medicine and the Department of Microbiology.

Edited by: T. P. Hatch