Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2014 Aug 1.
Published in final edited form as: Pathog Dis. 2013 Jul 10;68(3):88–95. doi: 10.1111/2049-632X.12052

Chlamydial infection of the gastrointestinal tract: a reservoir for persistent infection

Laxmi Yeruva 2, Nicole Spencer 1, Anne K Bowlin 1, Yin Wang 1, Roger G Rank 1,*
PMCID: PMC3751173  NIHMSID: NIHMS496918  PMID: 23843274

Abstract

The mechanism by which chlamydiae persist in vivo remains undefined; however, chlamydiae in most animals persist in the gastrointestinal tract (GI) and are transmitted via the fecal-oral route. Oral infection of mice with Chlamydia muridarum was previously shown to establish a long-term persistent infection in the GI tract. In this study, BALB/c, DBA/2 and C57Bl/6 mice, infected orally with C. muridarum, were infected in the cecum for as long as 100 days in the absence of pathology. The primary target tissue was the cecum although the large intestine was also infected in most animals. A strong serum IgG and cecal IgA antibody response developed. Lymphocyte proliferation assays to chlamydial antigen on mesenteric lymph node cells were positive by day 10 and peaked on days 15–21, but the response returned to baseline levels by 50 days, despite the ongoing presence of the organism in the cecum. Since studies have shown that women and men become infected orally with chlamydiae, we propose that the GI tract is a site of persistent infection and that immune down-regulation in the gut allows chlamydiae to persist indefinitely. As a result, women may become reinfected via contamination of the genital tract from the lower GI tract.

Introduction

One of the major questions associated with human chlamydial genital infection is why so many infections remain subclinical and apparently persist for long periods of time. A consequence of these long-term subclinical infections is that there may be continual tissue damage, especially if the organism ascends to the Fallopian tubes, leading to fibrosis, tubal obstruction and infertility. The mechanism(s) by which chlamydiae persist has been the subject of much controversy, and there is still no definitive answer to this question. The predominant hypothesis is that the organism enters into a non-replicating aberrant form in the genital tract, perhaps under the influence of gamma interferon (IFN-γ). However, with the exception of a single study with C. muridarum in the mouse genital tract very early in the infection, visual evidence for aberrant chlamydiae in the genital tract has not been found in vivo (Rank et al. 2011).

Nevertheless, persistent infections have been well-documented in sheep, pigs, cattle, and birds, and persistent infections are, in fact, felt to be a hallmark of chlamydial infection in those animals; so it stands to reason that persistent infections should also be present in humans. However, an often overlooked fact is that in virtually all Chlamydia species in animals, including birds, cattle, sheep, pigs, mice, and guinea pigs, chlamydiae target the gastrointestinal tract (GI) and are transmitted via the fecal-oral route. Thus, in all of these animals, the natural site of infection is the GI tract. Indeed, it was recognized decades ago that chlamydiae persisted in the GI tract for long periods of time and that “the infectious chain must be tightly linked to the infectious fecal excretions (Storz 1971).” Moreover, Storz observed over 45 years ago that infection persisted in the lower GI tract of sheep and even if animals had a high titer of antibody, they were still susceptible to infection in the gut (Storz 1964). Recently, Pospischil and colleagues published histopathologic and electron microscopic images of GI infection of pigs with C. suis and observed both normal and aberrant chlamydial forms (Pospischil et al. 2009). More importantly, natural infections with C. suis are often sub-clinical, and interestingly, no obvious inflammatory response was noted in any of the GI tissue sections.

Using the mouse model, Igietseme and colleagues demonstrated that C. muridarum can persist in the GI tract of mice for up to 260 days (Perry & Hughes 1999; Igietseme et al. 2001). Of interest was the complete lack of pathology in the GI tract of the infected mice over the entire time course. In contrast, chlamydial infection of the cervix and upper genital tract in mice and guinea pigs induces a strong inflammatory response and resolves in 3–4 weeks following onset of the adaptive immune response (Rank & Sanders 1992; Morrison & Morrison 2000). In fact, the GI tract would be an ideal site in which chlamydiae can persist similar to other gut microbiota because of a down-regulation of the host response. There is strong documentation that the immune response in the GI tract is actually down-regulated by specific bacteria (Sokol et al. 2008; Round et al. 2011). Chlamydiae may persist in the GI tract either by down-regulating pathologic pro-inflammatory immune responses themselves or by taking advantage of those mechanisms elicited by other commensal bacteria, thereby allowing the GI tract to serve as a reservoir for (re)infection of the genital tract.

Since GI infection is the norm in most animal species, it is very likely that men and women become infected in the GI tract as well, and there is certainly clinical evidence to support this (Jones et al. 1985; Bax et al. 2011). If indeed chlamydiae become persistent in the GI tract, then there is always the risk of reinoculation of the genital tract from organisms shed in the rectum; thus persistence in humans may be more closely related to the site of infection rather than an alternative metabolic form. In order to further understand the nature of the persistent infection in the GI tract, further information on the actual site of infection, the kinetics of the infection and the nature of the local immune response are required. Therefore, in this study, we have extended the studies published by Perry to characterize in greater detail the long-term infection of C. muridarum in the GI tract of the mouse with emphasis on the humoral and cell-mediated immune response.

Materials and Methods

Experimental animals

Six-week old C57Bl/6 mice, BALB /c and DBA/2 mice were obtained from Jackson Laboratories (Bar Harbor, ME) and Harlan-Sprague Dawley (Indianapolis, IN) and were housed in a barrier facility with a 12:12 light:dark cycle and provided food and water ad libitum.

Mice were infected orally with 3 × 106 inclusion forming units (IFU) of C. muridarum, suspended in 0.1 ml sucrose-phosphate-glutamate buffer (SPG) by gavage. C. muridarum (Nigg strain) was originally obtained from the American Type Culture Collection as a yolk sac preparation about 1977 and has been passaged continuously in this laboratory since that time, first in yolk sacs and then in tissue culture. Mice were also inoculated orally with 3 × 106 IFU of C. trachomatis serovar E originally obtained from the University of Washington. All protocols were approved by the Institutional Animals Care and Use Committee.

Chlamydial culture

In order to quantify the number of chlamydiae in GI tissue, the gut was removed and dissected into individual portions of the jejunum, ileum, caecum and large intestine. Each tissue was dissected longitudinally and the contents removed by washing with phosphate-buffered saline (PBS). The epithelium was gently scraped with a scalpel blade and deposited into a sterile Eppendorf tube containing two 4 mm glass beads and 1 ml of 2-sucrose-phosphate buffer transport medium with 0.1 mg gentamicin, 0.2 mg vancomycin, and 2.5 μg of Fungizone. The tubes were vortexed for one minute and then sonicated for one minute. After centrifugation to pellet the large cell debris, the supernatants were diluted with SPG and inoculated onto cell monolayers for the determination of IFU. The number of IFU was determined by culturing in either McCoy or HeLa cells according to standard protocol (Hough & Rank 1988). We found that removal of food for 12 hours prior to euthanasia and culture of cecal scrapings in HeLa cells increased the sensitivity of detection of chlamydiae in the cecum.

Proliferation assay

Mesenteric lymph node (MLN) and iliac lymph node (ILN) cells were collected in RPMI 1640 medium and minced into single cell suspensions. Because of the small size of the ILN, all ILN from a group of mice were pooled. The cells were put through a 70 μm cell strainer and washed with medium 3 times. The cells (2 × 105 per well) were added to a 96-well plate along with 5 μg of UV-inactivated gradient-purified C. muridarum elementary bodies. Concanavalin A (5 μg per well) was added to designated wells as a positive control. Each sample was done in triplicate. At 72 hours of incubation, 20 μl of alamar Blue (Invitrogen Corporation, Carlsbad, CA) was added to each well and the absorbance determined 24 hours later at 570 nm and 600 nm (Ahmed et al. 1994). The percent difference in reduction of alamar Blue by antigen-treated versus control untreated cells was calculated in order to determine the level of proliferation.

Assessment of antibody levels

Serum and cecal antibodies to chlamydiae were determined by ELISA. C. muridarum antigen was purified as previously described (Hough & Rank 1988), and serum or cecal contents extract were added to the plate with an initial dilution of 1:10. Goat horseradish peroxidase conjugated anti- mouse IgG antibody were obtained from Southern Biotechnology Associates (Birmingham, AL) and goat horseradish peroxidase conjugated anti-mouse IgA was obtained from Serotec (Raleigh, NC). The color reaction was developed with ABTS (Sigma, St. Louis, MO, Catalogue #A1888). In order to assess the level of local IgA antibody in cecal contents, the cecal material was collected in PBS in a weight/volume ratio normalized to the lowest weight cecal contents. The material was stored at -70 until it could be assayed. Prior to assay, the suspension was centrifuged to pellet debris and the supernatant used for the antibody assay.

Histopathology

Tissues were fixed directly in buffered formalin and were then prepared and stained with hematoxylin and eosin according to standard methodology. Chlamydial inclusions were directly visualized on tissue sections by immunohistochemistry. Briefly, sections were incubated with a monoclonal mouse anti-chlamydial lipopolysaccharide (LPS) antibody prepared from the clone EVI H1 (a kind gift of Dr. You-xun Zhang, Boston University) followed by reagents from a horseradish peroxidase-DAB kit (R&D).

Results

Course and site of GI infection

Initially, we wanted to determine the primary site of chlamydial infection in the GI tract over an extended period of time. Thus, we inoculated mice orally with 3 × 106 IFU of C. muridarum and on days 5, 10, 15, 25, 50, 75, and 100 days post-infection, euthanized 4–5 five mice at each time and processed the jejunum, ileum, cecum and large intestine for isolation of chlamydiae. We found that mice were routinely positive in the cecum at all time points (Table 1). On day 5, 3 of 5 mice were positive and on day 100, 4 of 5 mice were positive, but at all other times, 5 of 5 mice were positive in the cecum. The cecum clearly was the target tissue of GI infection. In contrast, the jejunum was rarely positive, only 2 animals on day 5 and 1 on day 75. Similarly, mice were only positive in the ileum on days 10–25 and not thereafter. There was a high level of infection in the large intestine early in the infection, and some mice were still positive on days 50 and 75, indicating that the large intestine may be a secondary site of infection or organisms were in the process of being excreted.

Table 1.

Isolation of C. muridarum from gastrointestinal tract following oral inoculation

Day After Infection Jejunum Ileum Cecum Large intestine
5 0/4* 0/5 3/5 3/5
10 2/3 1/1 4/4 4/4
15 0/4 2/5 5/5 5/5
25 0/5 1/5 5/5 4/5
50 0/2 0/2 5/5 2/4
75 1/5 0/5 5/5 1/5
100 ND** ND 4/5 ND
Open in a new tab
*

No. positive/No. tested

**

Not done

While Igietseme and Perry had isolated chlamydiae from the large intestine up to 260 days after infection (Perry & Hughes 1999; Igietseme et al. 2001), they had not characterized the course of the infection. Therefore, in three separate experiments, using three strains of mice, C57Bl/6, BALB/c, and DBA/2 mice were inoculated with 2 × 106 IFU of C. muridarum orally, and ceca from 5 mice of each strain were collected at various times after infection for isolation of chlamydiae. At every time point at which ceca were assessed for chlamydial infection, almost all animals were infected, regardless of strain (Table 2). At day 75, a total of 33 of 33 mice were still infected.

Table 2.

Number of mice isolation-positive in the cecum

Day After Infection C57Bl/6 BALB/c DBA/2
5 8/10* 5/5 5/5
10 12/14 8/10 5/5
15 15/15 10/10 5/5
21 10/10 10/10 5/5
25 5/5 ND ND
35 10/10 9/10 5/5
50 14/15 9/10 5/5
75 17/17 11/11 5/5
100 4/5 ND ND
Open in a new tab
*

No. positive/No. tested

**

Not done

Infection was established in the ceca of all three strains at day 5 and remained elevated through day 10 of BALB /c and DBA/2 mice but decreased in C57Bl/6 mice (Figure 1). In all strains of mice, the infection continued to decrease after day 10 until day 35, after which the levels of organisms were relatively unchanged through day 75. It is interesting to note that the course of infection in C57Bl/6 mice was significantly different from BALB/c and DBA/2 mice (p<0.002 and P<0.001 respectively according to a 2 factor [group and days] ANOVA), primarily being lower in the first 25 days of infection. In contrast to genital infection, the total number of organisms in the entire cecum was low, ranging from a median of 103 to 105 IFU. Nevertheless, the important finding was that the infection did not resolve in the GI tract as it does in the genital tract. Moreover, even though C57Bl/6 mice had lower levels of infection early in the course, all strains had persistent infection through day 75.

Figure 1.

Figure 1

Open in a new tab

Kinetics of infection in the cecum of C57Bl/6, BALB/c, and DBA/2 mice. Each point is the mean and standard deviation of 5 mice.

It was also important for the establishment of this model that we determine the infectious dose 50 (ID50) for GI infection. Groups of five C57Bl/6 mice each were inoculated orally with 102, 103, 104, 105, and 106 IFU and the IFU in the ceca determined at day 35 after inoculation. Because 4 of 5 mice were still positive at the 102 dose, the ID50 was considered to be less than 100 IFU.

While C. muridarum is a natural chlamydia of mice, we wanted to determine if we could also infect mice in the GI tract with C. trachomatis. Mice can be infected in the genital tract with C. trachomatis but the infection is far less intense than C. muridarum, and progesterone treatment is required for the infection to be established. Therefore, in two separate experiments, mice were inoculated orally with C. trachomatis, serovar E, and ceca were collected from five mice each on days 10 and 52 in one experiment and days 15 and 35 in the second experiment. In every animal, the cecum was negative for chlamydiae, indicating that C. trachomatis is unable to colonize the GI tract in mice.

Histopathology

Previously, Perry and coworkers reported that there were no detectable pathologic changes in the GI tract of mice infected orally with C. muridarum at any time after infection. We examined the ceca of 5 mice 15 days after infection and 2 mice 25 days after infection and also observed that the ceca were totally normal in appearance. We also stained sections by immunohistochemistry with antibody to chlamydial LPS in an effort to detect chlamydial inclusions and localize them within the cecum. Inclusions were very rare; usually just one on a section but it appeared that they were present in the proximal and distal portions of the cecum in areas where there were more convolutions in the epithelium (Figure 2). We did not observe inclusions in the middle areas of the cecum where the epithelium was thinner and less convoluted. As predicted by the overall histopathologic examination and the previous report (Igietseme et al. 2001), we did not see any inflammatory response associated with the inclusions.

Figure 2.

Figure 2

Open in a new tab

Localization of chlamydial inclusion in cecal epithelium 25 days after infection. Note the absence of an inflammatory response. The inset box is a higher magnification of the area containing the chlamydial inclusion.

Host immune response to oral infection

In order to assess the antibody immune response following GI infection, sera and cecal contents for systemic IgG and IgA antibody levels respectively were collected from 5–7 C57Bl/6 and BALB/c mice each on days 0, 10, 15, 21, 35, 50, and 75. MLN and ILN were collected from each animal for the assessment of T cell proliferation to chlamydial antigen. In addition, cecal tissue was collected for isolation of chlamydiae. All mice were indeed found to be positive for chlamydiae in the ceca.

When anti-chlamydial antibody was assessed in sera, IgG was first detected consistently on day 15 in both strains of mice and increased to high levels by day 25. IgG in C57Bl/6 mice continued to increase until the end of the experiment while IgG in BALB/c mice remained relatively level until the end of the experiment (Figure 3). The IgG response in C57Bl/6 mice was significantly higher than the BALB/c mice (P=0.004, 2 factor [group and days] ANOVA). Similarly, IgA antibody to chlamydial antigen was measured in the cecal contents and was first observed to be positive on day 15 after infection in both groups of mice and increased to a peak levels on day 50. By day 75, cecal IgA levels had decreased somewhat. Levels of IgA in C57Bl/6 mice were significantly lower than BALB/c mice (p=006, 2 factor [group and days] ANOVA) as a result of differences on days 15 and 35 (P=0.001 and p=0.003, Tukey analysis). No IgG antibody was detected in cecal contents.

Figure 3.

Figure 3

Open in a new tab

Kinetics of the serum and cecal antibody responses in C57Bl/6 and BALB/c mice infected orally with C. muridarum. Upper panel - Serum IgG antibody response to C. muridarum antigen. Each point is the mean and standard deviation of 5 mice. Lower panel – Cecal IgA antibody response to C. muridarum antigen. Each point is the mean and standard deviation of 5 mice.

Because the MLN are the draining lymph nodes of the GI tract, cell-mediated immunity was assessed by proliferation of MLN lymphocytes to chlamydial antigen (Figure 4). In both C57Bl/6 and BALB/c mice, MLN cells responded by day 10 and reached peak levels on days 25 in C57Bl/6 mice and 10–15 in BALB/c mice. Concanavalin A responses were always positive demonstrating viability of the cells and functionality of the assay (data not shown). It is interesting that after reaching a peak level relatively early in the infection course, the proliferative response began to decrease in both strains of mice and returned to pre-infection levels by day 50 even though all mice continued to be infected in the cecum. There was no significant difference between strains when analyzed with a two-way [group, days] ANOVA. We also assessed the proliferative response to chlamydial antigen by ILN lymphocytes (Figure 5). Because there were so few cells in the ILN, we pooled the nodes from all mice in a group. Nevertheless, the response in the ILN roughly paralleled that in the MLN, particularly with the return of the response to baseline levels by days 50 and 75. Thus, the data indicate that GI infection can induce a response in the lymph nodes draining the genital tract possibly by homing of T cells originally sensitized in the MLN.

Figure 4.

Figure 4

Open in a new tab

Kinetics of lymphocyte proliferation to C. muridarum antigen on MLN from orally-infected C57Bl/6 and BALB/c mice. Each point is the mean and standard deviation of 5 mice.

Figure 5.

Figure 5

Open in a new tab

Kinetics of lymphocyte proliferation to C. muridarum antigen on ILN from orally-infected C57Bl/6 and BALB/c mice. Each point represents pool of 5 mice.

Discussion

In this study, we have confirmed the observations by Igietseme and Perry that C. muridarum can colonize the gastrointestinal tract of mice indefinitely without causing any pathologic response and have extended those studies to more fully characterize the course of gastrointestinal infection and the resulting host response (Perry & Hughes 1999; Igietseme et al. 2001). Following oral inoculation, chlamydiae could be detected primarily in the cecum and large intestine, with peak levels about 10 days after infection. The level of infection decreased thereafter but animals were still infected 75–100 days after infection. The target sites were clearly the cecum and large intestine, although isolations in the large intestine declined after 10 days. The number of IFU was relatively consistent from days 35–75. It was also interesting that the ID50 was less than 100 IFU. This would indicate that chlamydial elementary bodies are relatively resistant to the acidity in the stomach. This also supports the likely fecal-oral route of transmission as so few organisms are required for infection.

It is not surprising that C. muridarum infection persists in the GI tract since in virtually all mammals and birds, the GI tract is the natural site of infection, and transmission is by the fecal-oral route. For instance, Storz and Thornely demonstrated that sheep were infected with chlamydiae in the cecum and continued to shed organisms for at least 4 years (Storz & Thornley 1966). It has always been assumed that the natural site of C. muridarum infection was the respiratory tract because it was originally isolated from that site. However, there is no evidence for horizontal transmission via aerosol (Karr 1943). In fact, Karr was only able to demonstrate horizontal transmission by either adding infected lung material to drinking water or adding mouse carcasses to cages of uninfected mice. Only then could she demonstrate chlamydiae in the lungs, suggesting oral transmission with aspiration to the lungs. Similarly, Cotter and coworkers housed uninfected mice with mice infected genitally with C. muridarum (Cotter et al. 1997). Naïve mice became positive in the mesenteric lymph nodes and at no other site, again suggesting oral transmission via grooming or coprophagy. Perry noted that mice infected intravaginally or intranasally also became positive in the GI tract (Perry & Hughes 1999). Similarly, we have observed that mice infected genitally and intranasally become positive in the cecum (unpublished data). In fact, in a recent experiment, 15 of 20 mice infected intranasally were infected in the cecum at day 35 after infection (unpublished data). Thus, just as in other species, the natural site of chlamydial infection in the mouse would appear to be the GI tract, and in particular, the cecum and large intestine.

To determine if there was any difference in the infection course in the GI tract related to the strain of mouse infected, we evaluated the infection course in BALB/c, C57Bl/6, and DBA/2 mice. Differences were noted between BALB/c and C57Bl/6 and between C57Bl/6 and DBA/2 but overall, the kinetics of the infection in each strain was essentially the same. At this point, it is not clear whether the strain-dependent differences represent any true impact on the nature of GI infection in mice. However, it is interesting to note that the BALB/c mice have a higher level of infection than the C57Bl/6 mice in the first 3 weeks of infection. We had previously observed that BALB/c mice infected genitally with C. muridarum also had longer infections with more upper tract pathology than C57Bl/6 mice (Darville, T., Rank, R.G., unpublished data).

Although the strain of mouse did not seem to influence the ability of C. muridarum to infect the GI tract, there appeared to be a definite chlamydial species specificity. In two separate experiments, we tried to infect mice orally with C. trachomatis, serovar E, and were unsuccessful. This is not surprising, since C. trachomatis is not a natural chlamydia of mice and does not produce as productive infection in the genital tract as does C. muridarum.

As first observed by Igietseme and Perry (Igietseme et al. 2001), we also found that there was no pathologic response associated with infection in the cecum; therefore, it is apparent that there is either an absence of or down-regulation of the local innate and adaptive immune response. It is possible that there may be an early innate response, but we did not evaluate tissues at the very early stage of the infection. That the host does indeed respond to GI infection was noted by the presence of a strong local and systemic antibody response as well as a positive proliferation response in the MLN and ILN. Moreover, previous studies with oral immunization with viable organisms in both mouse and guinea pig models have demonstrated a substantial humoral and cell-mediated protective response to genital tract challenge (Nichols et al. 1978; Rank et al. 1990; Kelly et al. 1996). The absence of an inflammatory response in conjunction with clear evidence for active replication of chlamydiae as evidenced by successful isolation could be the result of numerous well-documented down-regulatory mechanisms elicited by other gut microbiota (Round & Mazmanian 2009). For example Bacterioides fragilis, a gut commensal, activates the TLR2 pathway which elicits a Treg cell response to down-regulate the host response in the GI tract (Round et al. 2011). Bifidobacterium infantis has also been found to induce Treg cells and inhibit NF-κB activation (O’Mahony et al. 2008).

That the infection in the GI tract is elevated initially and then declines after two weeks suggests that a protective immune response is indeed elicited and is effective in reducing the number of organisms. The decrease in the level of infection corresponds to an increasing local IgA response in the cecum, serum IgG response, and cell-mediated immune response in the draining MLN. However, after 35 days, the infection decreases to a low but steady-state level, concomitant with a return of the T cell proliferative response in the MLN to baseline levels. IgA is still present but also begins to decrease by day 75. In the mouse genital and respiratory models, a Th1 response is essential for final clearance of the infection (Coalson et al. 1987; Igietseme et al. 1993; Magee et al. 1993; Cain & Rank 1995), so the absence of a local T cell response in the GI tract in the latter stage of the infection may be preventing the organism from being eliminated. Nevertheless, IgA antibody is still present throughout the period studied, so it may be responsible for holding the infection in check at a low level but is unable to effect complete resolution. In the guinea pig genital tract model, antibody is necessary for both resolution of and resistance to reinfection but requires cell-mediated immunity to eliminate the infection (Rank et al. 1979; Rank & Barron 1983). Antibody is also important for resistance to reinfection in the mouse model (Morrison & Morrison 2005).

The question then arises as to why the T cell response decreases to pre-infection levels even in the presence of organisms. One explanation is that the number of chlamydiae has been reduced to very low levels by day 35 and thereafter, between 103 – 104 IFU per gram of cecum which translates into about 102–103 IFU for the entire cecum. It is quite possible that as the organism decreases in number, there is less antigen available to stimulate the immune response, and a level may be reached which is sub-immunogenic. The inflammatory response is likely down-regulated through mechanisms associated with other microbiota, so in the absence of inflammatory cells and a local adaptive immune response, the organisms are able to persist indefinitely. Alternatively, but not necessarily exclusively, the lack of an inflammatory response indicates that there are either no or only minimal pro-inflammatory cytokines such as TNF-α being produced; so consequently, there is no up-regulation of addressins such as VCAM-1 and MadCAM-1, without which T cells, macrophages, and PMNs cannot home to the site.

Nevertheless, from the perspective of Chlamydia, the GI tract is an ideal location for chlamydiae because they can continue to replicate, and some EBs will be liberated into the lumen of the gut where they are excreted and can be transmitted to new hosts by the fecal-oral route. Moreover, there is a high turnover rate of epithelial cells in the gut, providing an ample number of new target cells for chlamydiae.

Regardless of the mechanisms by which chlamydiae survive in the local GI milieu, oral infection clearly activates an adaptive immune response that may manifest protection in other mucosal sites. In both mouse and guinea pig models, oral infection can elicit a protective response in the genital tract resulting in a shorter course of infection with lower peak infection levels (Nichols et al. 1978; Rank et al. 1990; Kelly et al. 1996). We noted many years ago that serum antibodies to all chlamydial antigens actually increased over time and that antibody to higher molecular weight proteins appeared consistently 90–100 days after infection (Ramsey et al. 1989). At that time, we proposed that the infection may be persisting at some other site. It now seems very likely that all of the mice had developed GI infections and that the constant exposure to chlamydial antigens elicited the more pronounced serum antibody response over time.

That chlamydiae can persist in the GI tract in virtually all animals begs the question as to why chlamydiae could not persist in the GI tract of humans as well. There is substantial clinical data to indicate that women become infected in the GI tract. Jones and colleagues isolated chlamydiae from the pharynx of 3.2% and rectum of 5.2% of women attending a sexually transmitted disease clinic (Jones et al. 1985). Interestingly, they found an incidence of 5.8% rectal positive cultures in women with no history of anal intercourse. In addition, they observed that 11% of women with genital infection and 17% of women with pharyngeal infection also had positive rectal cultures and that there was no correlation between women with positive rectal cultures and admitted anal intercourse. In a recent study by Bax and colleagues, 65.5% of women were found to be positive for Chlamydia and when assessed by cervical and rectal samples were positive in both sites (Bax et al. 2011). Five patients of 177 tested were only positive in the rectum. However, no specific information was given regarding sexual practices. While it is not uncommon to find women who are rectal positive for chlamydiae, the only evidence for long term infection was a prospective study by Bell and colleagues in which they obtained cultures from various anatomic sites of infants born to Chlamydia-infected mothers (Bell et al. 1992). The infants were undoubtedly infected at birth, and they found that 6 of 6 infants infected in the rectum remained positive in the rectum for at least 383 days. That chlamydiae persist in the GI tract of most animals species and have been documented to persist in the GI tract of mice for extended periods of time without causing pathology, suggests that it is entirely feasible that this could occur in humans as well.

If indeed this is the case, then the GI tract could be a reservoir for chlamydiae which would then be available to reinfect the genital tract. It is not difficult to conceive that women can become infected in the GI tract even in the absence of anal intercourse via fluid contamination of the rectum from the vagina or through oral sexual activity. Studies in all animal models demonstrate that genital infection resolves within several weeks through the development of a Th1 cell-mediated immune response (Rank 2006). However, because T cells do not remain in the genital tract following recovery, the individual becomes susceptible to reinfection (Igietseme & Rank 1991). Clearly, most reinfections likely result from repeated sexual activity with infected partners; however, it is quite possible that contamination of the genital tract from the rectum, may occur, providing an opportunity for chlamydiae in the GI tract to “reinfect” the genital tract. This is a reasonable scenario as women commonly develop urogenital infections with Escherichia coli whereas men do not. In mice, Perry and Hughes observed that orally infected female mice could develop infection in the genital tract (Perry & Hughes 1999). Therefore we suggest that persistence in chlamydial infections is associated with reseeding of the genital tract from chlamydiae colonizing and living as commensals in the GI tract.

The observation that long-term GI infection persists even in the presence of an antibody response, both local and systemic, has important implications for a chlamydial vaccine. While a vaccine may be effective in preventing or reducing genital chlamydial infection or disease, it is unlikely to eliminate chlamydiae from the GI tract. Thus, the subject would still maintain a persistent infection in the GI tract which could be spread to other individuals or potentially reinfect the genital tract. On the other hand, it is possible that persistence of the organism in the GI tract may provide a booster effect for protection in the genital tract or provide a source of T cells capable of homing to the genital tract where they could either provide a protective role or exacerbate genital tract pathology. We have reported previously that T cells homing to the genital tract bear the homing receptor, α4β7, the same homing receptors as on T cells homing to the GI tract and that MadCAM-1, the ligand for α4β7, is present in both the genital tract and the GI tract (Kelly & Rank 1997). Furthermore, if the chlamydiae in the GI tract are maintained at a low level through various down-regulatory mechanisms, disruption of these homeostatic mechanisms by antibiotics, diet, or any factor disrupting the GI microbiota could possibly increase the number of chlamydiae present and increase the likelihood for reinfection of the genital tract. Therefore, it will be critical that any studies investigating the effectiveness of a chlamydial vaccine also investigate the impact on chlamydial colonization of the GI tract.

Acknowledgments

This research was supported, in part, by Arkansas Children’s Hospital Research Institute and the Arkansas Biosciences Institute, the Marion B. Lyon New Scientist Development Award (VLY), and grant U19 AI084044 from the NIAID, NIH.

Reference List

  1. Ahmed SA, Gogal RM, Jr, Walsh JE. A new rapid and simple non-radioactive assay to monitor and determine the proliferation of lymphocytes: an alternative to [3H]thymidine incorporation assay. J Immunol Methods. 1994;170:211–224. doi: 10.1016/0022-1759(94)90396-4. [DOI] [PubMed] [Google Scholar]
  2. Bax CJ, Quint KD, Peters RPH, Ouburg S, Oostvogel PM, Mutsaers JAEM, Doerr PJ, Schmidt S, Jansen C, van Leeuwen AP, Quint WGV, Trimbos JB, Meijer CJLM, Morre SA. Analyses of multiple-site and concurrent Chlamydia trachomatis serovar infections, and serovar tissue tropism for urogenital versus rectal specimens in male and female patients. Sexually Transmitted Infections. 2011;87:503–507. doi: 10.1136/sti.2010.048173. [DOI] [PubMed] [Google Scholar]
  3. Bell TA, Stamm WE, Wang S, Kuo C, Holmes KK, Grayston JT. Chronic Chlamydia trachomatis infections in infants. JAMA. 1992;267:400–402. [PubMed] [Google Scholar]
  4. Cain TK, Rank RG. Local Th1-like responses are induced by intravaginal infection of mice with the mouse pneumonitis biovar of Chlamydia trachomatis. Infection and Immunity. 1995;63:1784–1789. doi: 10.1128/iai.63.5.1784-1789.1995. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Coalson JJ, Winter VT, Bass LB, Schachter J, Grubbs BG, Williams DM. Chlamydia trachomatis pneumonia in the immune, athymic and normal BALB mouse. Br J Exp Pathol. 1987;68:399–412. [PMC free article] [PubMed] [Google Scholar]
  6. Cotter TW, Ramsey KH, Miranpuri GS, Poulsen CE, Byrne GI. Dissemination of Chlamydia trachomatis chronic genital tract infection in gamma interferon gene knockout mice. Infection and Immunity. 1997;65:2145–2152. doi: 10.1128/iai.65.6.2145-2152.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Hough AJ, Jr, Rank RG. Induction of arthritis in C57Bl/6 mice by chlamydial antigen: Effect of prior immunization or infection. American Journal of Pathology. 1988;130:163–172. [PMC free article] [PubMed] [Google Scholar]
  8. Igietseme JU, Portis JL, Perry LL. Inflammation and clearance of Chlamydia trachomatis in enteric and nonenteric mucosae. Infection and Immunity. 2001;69:1832–1840. doi: 10.1128/IAI.69.3.1832-1840.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Igietseme JU, Ramsey KH, Magee DM, Williams DM, Kincy TJ, Rank RG. Resolution of murine chlamydial genital infection by the adoptive transfer of a biovar-specific TH1 lymphocyte clone. Regional Immunology. 1993;5:317–324. [PubMed] [Google Scholar]
  10. Igietseme JU, Rank RG. Susceptibility to reinfection after a primary chlamydial genital infection is associated with a decrease of antigen-specific T cells in the genital tract. Infection and Immunity. 1991;59:1346–1351. doi: 10.1128/iai.59.4.1346-1351.1991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Johansson ME, Phillipson M, Petersson J, Velcich A, Holm L, Hansson GC. The inner of the two Muc2 mucin-dependent mucus layers in colon is devoid of bacteria. Proc Natl Acad Sci USA. 2008;105:15064–15069. doi: 10.1073/pnas.0803124105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Jones RB, Rabinovitch RA, Katz BP, Batteiger BE, Quinn TS, Terho P, Lapworth MA. Chlamydia trachomatis in the pharynx and rectum of heterosexual patients at risk for genital infection. Annals of Internal Medicine. 1985;102:757–762. doi: 10.7326/0003-4819-102-6-757. [DOI] [PubMed] [Google Scholar]
  13. Karr HV. Study of a latent pneumotropic virus of mice. J Infect Dis. 1943;72:108–116. [Google Scholar]
  14. Kelly KA, Rank RG. Identification of homing receptors that mediate the recruitment of CD4 T cells to the genital tract following intravaginal infection with Chlamydia trachomatis. Infection and Immunity. 1997;65:5198–5208. doi: 10.1128/iai.65.12.5198-5208.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Kelly KA, Robinson EA, Rank RG. Initial route of antigen administration alters the T-cell cytokine profile produced in response to the mouse pneumonitis biovar of Chlamydia trachomatis following genital infection. Infect Immun. 1996;64:4976–4983. doi: 10.1128/iai.64.12.4976-4983.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Magee DM, Igietseme JU, Smith JG, Bleicker CA, Grubbs BG, Schachter J, Rank RG, Williams DM. Chlamydia trachomatis pneumonia in the severe combined immunodeficiency (SCID) mouse. Regional Immunology. 1993;5:305–311. [PubMed] [Google Scholar]
  17. Morrison SG, Morrison RP. In situ analysis of the evolution of the primary immune response in murine Chlamydia trachomatis genital tract infection. Infection and Immunity. 2000;68:2870–2879. doi: 10.1128/iai.68.5.2870-2879.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Morrison SG, Morrison RP. A predominant role for antibody in acquired immunity to chlamydial genital tract reinfection. The Journal of Immunology. 2005;175:7536–7542. doi: 10.4049/jimmunol.175.11.7536. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Nichols RL, Murray ES, Nisson PE. Use of enteric vaccines in protection against chlamydial infections of the genital tract and the eye of guinea pigs. J Infect Dis. 1978;138:742–746. doi: 10.1093/infdis/138.6.742. [DOI] [PubMed] [Google Scholar]
  20. O’Mahony C, Scully P, O’Mahony D, Murphy S, O’Brien F, Lyons A, Sherlock G, MacSharry J, Kiely B, Shanahan F, O’Mahony L. Commensal-induced regulatory T cells mediate protection against pathogen-stimulated NF-kappaB activation. PLoS Pathog. 2008;4:e1000112. doi: 10.1371/journal.ppat.1000112. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Perry LL, Hughes S. Chlamydial colonization of multiple mucosae following infection by any mucosal route. Infection and Immunity. 1999;67:3686–3689. doi: 10.1128/iai.67.7.3686-3689.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Pospischil A, Borel N, Chowdhury EH, Guscetti F. Aberrant chlamydial developmental forms in the gastrointestinal tract of pigs spontaneously and experimentally infected with Chlamydia suis. Vet Microbiol. 2009;135:147–156. doi: 10.1016/j.vetmic.2008.09.035. [DOI] [PubMed] [Google Scholar]
  23. Ramsey KH, Newhall WJ, Rank RG. Humoral immune response to chlamydial genital infection of mice with the agent of mouse pneumonitis. Infection and Immunity. 1989;57:2441–2446. doi: 10.1128/iai.57.8.2441-2446.1989. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Rank RG. The role of the CD4 T cell in the host response to Chlamydia. In: Bavoil PM, Wyrick PB, editors. Chlamydia: Genomics and Pathogenesis. Horizon Bioscience; Norfolk, UK: 2006. pp. 365–380. [Google Scholar]
  25. Rank RG, Barron AL. Humoral immune response in acquired immunity to chlamydial genital infection of female guinea pigs. Infection and Immunity. 1983;39:463–465. doi: 10.1128/iai.39.1.463-465.1983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Rank RG, Batteiger BE, Soderberg LSF. Immunization against chlamydial genital infection in guinea pigs with UV-inactivated and viable chlamydiae administered by different routes. Infection and Immunity. 1990;58:2599–2605. doi: 10.1128/iai.58.8.2599-2605.1990. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Rank RG, Sanders MM. Pathogenesis of endometritis and salpingitis in a guinea pig model of chlamydial genital infection. American Journal of Pathology. 1992;140:927–936. [PMC free article] [PubMed] [Google Scholar]
  28. Rank RG, White HJ, Barron AL. Humoral immunity in the resolution of genital infection in female guinea pigs infected with the agent of guinea pig inclusion conjunctivitis. Infection and Immunity. 1979;26:573–579. doi: 10.1128/iai.26.2.573-579.1979. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Rank RG, Whittimore J, Bowlin AK, Wyrick PB. In vivo ultrastructural analysis of the intimate relationship between polymorphonuclear leukocytes and the chlamydial developmental cycle. Infection & Immunity. 2011;79:3291–3301. doi: 10.1128/IAI.00200-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Round JL, Lee SM, Li J, Tran G, Jabri B, Chatila TA, Mazmanian SK. The Toll-like receptor 2 pathway establishes colonization by a commensal of the human microbiota. Science. 2011;332:974–977. doi: 10.1126/science.1206095. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Round JL, Mazmanian SK. The gut microbiota shapes intestinal immune responses during health and disease. Nat Rev Immunol. 2009;9:313–323. doi: 10.1038/nri2515. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Sokol H, Pigneur B, Watterlot L, Lakhdari O, Bermudez-Humaran LG, Gratadoux JJ, Blugeon S, Bridonneau C, Furet JP, Corthier G, Grangette C, Vasquez N, Pochart P, Trugnan G, Thomas G, Blottiere HM, Dore J, Marteau P, Seksik P, Langella P. Faecalibacterium prausnitzii is an anti-inflammatory commensal bacterium identified by gut microbiota analysis of Crohn disease patients. Proc Natl Acad Sci USA. 2008;105:16731–16736. doi: 10.1073/pnas.0804812105. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Storz J. Uber eine naturliche infektion eines meerschweinchenbestandes mit einem erreger aus der psittakose-lymphogranuloma gruppe. Zentralbl Bakteriol[A] 1964;193:432–446. [PubMed] [Google Scholar]
  34. Storz J. Intestinal infections of ruminants. In: Storz J, editor. Chlamydia and Chlamydia-induced Diseases. Charles C. Thomas; Springfield, IL: 1971. pp. 146–154. [Google Scholar]
  35. Storz J, Thornley WR. Serological and etiological studies on the intestinal psittacosislymphogranuloma infection in sheep. Zentralbl Veterinarmed B. 1966;13:14–24. [PubMed] [Google Scholar]

RESOURCES