ABSTRACT
The cryptic plasmid is important for chlamydial colonization in the gastrointestinal tract. We used a combination of intragastric, intrajejunal, and intracolon inoculations to reveal the impact of the plasmid on chlamydial colonization in distinct regions of gastrointestinal tract. Following an intragastric inoculation, the plasmid significantly improved chlamydial colonization. At the tissue level, plasmid-positive Chlamydia produced infectious progenies throughout gastrointestinal tract. However, to our surprise, plasmid-deficient Chlamydia failed to produce infectious progenies in small intestine, although infectious progenies were eventually detected in large intestine, indicating a critical role of the plasmid in chlamydial differentiation into infectious particles in small intestine. The noninfectious status may represent persistent infection, since Chlamydia genomes proliferated in the same tissues. Following an intrajejunal inoculation that bypasses the gastric barrier, plasmid-deficient Chlamydia produced infectious progenies in small intestine but was 530-fold less infectious than plasmid-positive Chlamydia, suggesting that (i) the noninfectious status developed after intragastric inoculation might be induced by a combination of gastric and intestinal effectors and (ii) chlamydial colonization in small intestine was highly dependent on plasmid. Finally, following an intracolon inoculation, the dependence of chlamydial colonization on plasmid increased over time. Thus, we have demonstrated that the plasmid may be able to improve chlamydial fitness in different gut regions via different mechanisms, which has laid a foundation to further reveal the specific mechanisms.
INTRODUCTION
Sexually transmitted Chlamydia trachomatis may ascend to the upper genital tract to cause pathology, leading to hydrosalpinx and tubal infertility (1–3). However, the pathogenic mechanisms remain unclear. Chlamydia muridarum infection in the mouse genital tract has been used extensively for investigating the mechanisms of C. trachomatis pathogenesis (4–9), because mice intravaginally inoculated with C. muridarum consistently develop hydrosalpinx and infertility (10–12). Using this murine model, both chlamydial (13–17) and host (4, 8, 18–24) factors that affect C. muridarum induction of hydrosalpinx have been identified. The cryptic plasmid, termed pMoPn/pCM (25, 26), is a known pathogenic determinant of C. muridarum in the mouse upper genital tract (13, 16, 17, 27, 28). The plasmid codes 8 putative proteins designated plasmid-encoded glycoproteins 1 to 8 (pGP1 to -8) (25, 28, 29). While pGP3 may directly exert virulence functions (13), pGP4 (30, 31) and pGP5 (32) regulate other chlamydial genes.
C. trachomatis is also frequently detected in the human gastrointestinal (GI) tract (33–36). Interestingly, when C. muridarum was introduced to multiple mouse mucosae, it readily colonized the GI tract (37). The genital C. muridarum selectively spread to (38) and established long-lasting colonization in (38–40) the GI tract lumen. The C. muridarum spreading is likely via a hematogenous route (41). Although C. muridarum colonization in the GI tract is long lasting, it is nonpathological in the GI tract (42, 43). However, it was hypothesized that the GI C. muridarum might induce immune responses that may contribute to clinically relevant pathologies in the female reproductive tract (44). Nevertheless, C. muridarum is well adapted to the GI tract environment. Various C. muridarum virulence determinants identified in the genital tract were found to be important for C. muridarum colonization in the GI tract (44–49). Thus, evaluating the roles of chlamydial factors in C. muridarum colonization in the GI tract may provide biologically relevant information for understanding chlamydial pathogenic mechanisms in the genital tract.
We recently showed that plasmid-deficient C. muridarum exhibits significantly delayed/reduced colonization in mouse GI tract (45) and that C. muridarum lacking the plasmid-encoded pGP3 is rapidly cleared in the stomach (47, 49), indicating a critical role of the plasmid in protecting C. muridarum against gastric killing. However, it is still unknown whether and how the plasmid contributes to C. muridarum colonization in the remaining regions of the GI tract. The present study was designed to evaluate the role of plasmid in regulating chlamydial fitness in different regions of GI tract by using a combination of intragastric, intrajejunal, and intracolon inoculations. Following an intragastric inoculation, although plasmid-positive Chlamydia produced infectious progenies throughout the gastrointestinal tract, plasmid-deficient Chlamydia failed to produce any detectable infectious progenies in small intestine. To exclude an effect from the gastric barrier, we further used an intrajejunal inoculation to deliver chlamydial organisms into the lumen of small intestine and found that plasmid-deficient Chlamydia was able to produce infectious progenies but was 530-fold less infectious than plasmid-positive Chlamydia. These observations suggest that the noninfectious status developed after intragastric inoculation may be induced by a combination of gastric and intestinal effectors and that Chlamydia colonization in small intestine is highly dependent on plasmid. Following an intracolon inoculation, the plasmid also improved Chlamydia colonization, and the plasmid-dependent improvement became more obvious with a prolonged colonization time course. These observations have led us to propose that Chlamydia may have acquired the plasmid for improving its fitness in different regions of the GI tract via different mechanisms, since different regions of the GI tract are equipped with distinct host defense effector mechanisms (50–52).
RESULTS
Lack of plasmid significantly compromises chlamydial ability to colonize mouse gastrointestinal tract.We previously demonstrated that the cryptic plasmid, a known genital tract pathogenic virulence determinant, is more important for promoting chlamydial colonization in the gastrointestinal tract than in the genital tract (45, 47). Here, we further quantitatively titrated the role of the plasmid in promoting chlamydial colonization in the GI tract following an intragastric inoculation at various inoculation doses (Fig. 1). At an inoculation dose of 1,000 inclusion-forming units (IFU) per mouse, plasmid-positive Chlamydia established colonization in the GI tracts of 100% of the mice by day 7 and maintained a steady level of colonization thereafter, which is consistent with what we showed previously (42). However, plasmid-deficient Chlamydia was able to do so only at the inoculation dose of 1 × 105 IFU per mouse or higher. Moreover, even at high inoculation doses, colonization by plasmid-deficient Chlamydia was significantly delayed, and live organisms were only detectable in the rectal swabs by day 14. Thus, the cryptic plasmid improved chlamydial colonization in the GI tract by >100-fold.
Comparison of live organism shedding between mice infected intragastrically with Chlamydia muridarum with or without plasmid. Groups of C57BL/6J (C57) mice (n = 4 to 6) inoculated intragastrically with C. muridarum with (plasmid positive) (a to d) or without (plasmid deficient) (e to h) plasmid were monitored for live chlamydial organism shedding in rectal swabs on days 3 and 7 and weekly thereafter, as indicated along the x axis. Different inoculation doses ranging from 103 to 106 inclusion forming units (IFU) per mouse were used for different groups of mice as indicated on the right, while the live organisms recovered from rectal swabs are expressed as log10 IFU as shown along the left y axis.
Plasmid is critical for promoting chlamydial colonization in the small intestine.To evaluate the role of the plasmid in improving chlamydial fitness in the small intestine, we first compared the yields of infectious organisms harvested from different GI tract tissues of mice intragastrically inoculated with Chlamydia with or without plasmid (Fig. 2). Infectious plasmid-positive Chlamydia was readily detected in all tissues throughout the GI tract but was gradually restricted to the large intestinal tissues by day 28. This tissue distribution pattern is consistent with what we reported previously (38). However, to our surprise, no infectious plasmid-deficient Chlamydia was detected in the small intestine during the entire course of the experiment, although chlamydial genomes were detectable throughout the GI tract (Fig. 3). The chlamydial genome copies increased over time, suggesting that chlamydial genomes were able to proliferate in the small intestine. By day 14 after inoculation (when live plasmid-deficient chlamydial organisms were detected in the rectal swabs) (see Fig. 1), the large intestine became positive for infectious organisms, and the number of large intestinal infectious organisms increased over time (Fig. 2), indicating that plasmid-deficient chlamydial organisms were able to produce infectious progenies in large intestine. Thus, in the absence of plasmid, chlamydial organisms were inhibited from producing infectious progenies only in small intestine. The question is, what host effectors are responsible for the suppression?
Comparison of live organism recovery from gastrointestinal tissues between mice inoculated intragastrically with Chlamydia muridarum with or without plasmid. Groups of C57 mice (n = 3 to 5) intragastrically inoculated with 1 × 105 IFU of Chlamydia muridarum with (plasmid positive) (a to e) or without (plasmid deficient) (f to j) plasmid were sacrificed to quantitate live chlamydial organism recovery in different regions of gastrointestinal tract as shown along the x axis on days 3 and 7 and weekly thereafter, as indicated on the right. The live organisms recovered from different tissues are expressed as log10 IFU as shown along the left y axis.
Chlamydial genome copies recovered from gastrointestinal tissues of mice inoculated intragastrically with plasmid-deficient Chlamydia. Groups of C57 mice (n = 3 to 5) intragastrically inoculated with 1 × 105 IFU of plasmid-deficient Chlamydia muridarum (as described in the legend for Fig. 2) were sacrificed to quantitate chlamydial genome copies in different regions of gastrointestinal tract as shown along the x axis on days 3 (a) and 7 (b) and weekly thereafter (c to e), as indicated on the right. The genome copies recovered from different tissues are expressed as log10 genomes per tissue as shown along the y axis.
To evaluate the role of small intestinal effectors in regulating plasmid-free Chlamydia colonization in the small intestine without interference from gastric effectors, we used an intrajejunal inoculation to bypass the gastric barrier (Fig. 4). We found that following intrajejunal inoculation, live plasmid-deficient chlamydial organisms were recovered from the small intestine, suggesting that the small intestinal noninfectious status of the plasmid-deficient chlamydial organisms developed following intragastric inoculation was caused by a combination of gastric and small effectors. Nevertheless, following intrajejunal inoculation, the plasmid deficiency increased the 50% infective dose (ID50) by ∼530-fold. Careful analyses further revealed that plasmid-positive Chlamydia maintained similar ID50s in both jejunum and colon following an intrajejunal inoculation, suggesting that most live plasmid-positive organisms recovered from jejunum tissue may have spread to the large intestine. However, plasmid-deficient Chlamydia increased its ID50 by ∼3-fold in the colon, implying that only one-third of the live plasmid-deficient organisms successfully spread from jejunum to the colon.
Comparison of live organism recovery from gastrointestinal tissues between mice infected via intrajejunal inoculation with Chlamydia muridarum with or without plasmid. Groups of C57 mice (n = 3 to 5) infected via intrajejunal inoculation with different doses (as shown in individual panels) of C. muridarum with (plasmid positive) (a to d) or without (plasmid free) (e to h) plasmid were sacrificed to quantitate live chlamydial organism recoveries in different regions of gastrointestinal tract as shown along the x axis on day 3 after intrajejunal inoculation. The live organisms recovered from different tissues are expressed as log10 IFU as shown along the left y axis. The doses required for infecting 50% of mice (ID50) were calculated for samples collected from jejunum and colon and are listed at the bottom (below the corresponding samples).
Plasmid promotes chlamydial adaptation in large intestine.To evaluate the role of the plasmid in chlamydial colonization in the colon, we used an intracolon inoculation (Fig. 5). We found that the infectious yields of plasmid-positive Chlamydia were significantly higher than those of plasmid-deficient Chlamydia, suggesting that plasmid may also promote chlamydial colonization in the large intestine. We further observed that upon intracolon inoculation, plasmid-positive Chlamydia was able to rapidly establish stable colonization starting from day 3, while plasmid-deficient Chlamydia started with a very low titer of infectious particles on day 3 and a very high titer on day 7 followed by a rapid decline in titer on day 14. The titers became relatively stable thereafter. These observations suggest that the plasmid may speed up chlamydial adaptation to the colonic environment. Nevertheless, despite these differences, both types of organisms maintained long-lasting colonization, up to 63 days. To further obtain quantitative data, we titrated the live organism recoveries from mice intracolonically inoculated with different amounts of chlamydial organisms (Fig. 6). We found that intracolon inoculation of 100 IFU was sufficient for plasmid-positive Chlamydia to establish stable colonization, while 1,000 IFU was required for plasmid-deficient Chlamydia to do so. More interestingly, plasmid-positive Chlamydia reduced its ID50s by ∼20-fold over time after intracolon inoculation (there was a transient ID50 reduction by ∼70-fold on day 7 only), suggesting that plasmid-positive Chlamydia is able to improve its colonization over time by adapting to the large intestine environment. However, plasmid-deficient Chlamydia maintained similar levels of ID50s throughout the time course despite a transient reduction on day 7, suggesting that in the absence of the plasmid, chlamydial organisms were unable to improve their colonization in the large intestine over time.
Comparison of live organism shedding between mice infected via intracolon inoculation with Chlamydia muridarum with or without plasmid. Groups of C57 mice (n = 4 to 6) inoculated via intracolon inoculation with 1 × 105 IFU of C. muridarum with (plasmid positive) (a) or without (plasmid deficient) (b) plasmid were monitored for live chlamydial organism shedding in rectal swabs on days 3 and 7 and weekly thereafter, as indicated along x axis. The live organisms recovered from rectal swabs are expressed as log10 IFU, as shown along the y axis. The level of colonization by plasmid-deficient Chlamydia was significantly lower than that of plasmid-positive Chlamydia (area under curve, Wilcoxon rank sum, P < 0.05).
Comparison of live organism recovery from rectal swabs between mice infected via intracolon inoculation with Chlamydia muridarum with or without plasmid. Groups of C57 mice (n = 3 to 5) infected via intracolon inoculation with different doses (as shown in individual panels) of C. muridarum with (plasmid positive) (a to d) or without (plasmid deficient) (e to h) plasmid were monitored for live chlamydial organism recovery in rectal swabs on days 3 and 7 and weekly thereafter after intracolon inoculation (as shown along x axis). The live organisms recovered are expressed as log10 IFU, as shown along the y axis. The doses required for infecting 50% of the mice (ID50) were calculated for samples collected at individual time points and are listed at the bottom (below the corresponding time points).
DISCUSSION
Chlamydia is frequently detected in the GI tracts of both animals and humans (33–36, 38, 53–55). However, the mechanisms by which Chlamydia colonizes the gut remain unknown. Although C. muridarum plasmid is a known pathogenic determinant in the genital tract, it is more important for C. muridarum to colonize GI tract than genital tract (45), suggesting that the plasmid might have been acquired by C. muridarum to improve its fitness in the GI tract. Here, we have used C. muridarum colonization in the mouse GI tract as a model to reveal the roles of the cryptic plasmid in promoting chlamydial colonization in distinct regions of the GI tract. By using a combination of intragastric, intrajejunal, and intracolon inoculations, we have demonstrated a critical role of the plasmid in improving chlamydial colonization in multiple regions of the GI tract. First, following an intragastric inoculation, the plasmid improved chlamydial colonization efficiency by >100-fold, since it took an inoculum dose of 105 IFU of plasmid-deficient Chlamydia to achieve stable colonization in the GI tract, while 1,000 IFU of plasmid-positive Chlamydia was able to do so. Second, when live organisms from individual mouse GI tract tissues were compared, we found that no live plasmid-deficient chlamydial organisms were recovered from small intestine, although live organisms were detected in the large intestines of the same mice on day 14 after intragastric inoculation, suggesting a requirement of the plasmid for chlamydial organisms to produce infectious progenies in the small intestine. Third, following an intrajejunal inoculation, infectious particles of plasmid-deficient Chlamydia were recovered from the small intestine but were ∼530-fold less infectious than those of plasmid-positive Chlamydia, indicating a critical role of the plasmid in chlamydial resistance to small intestinal effectors. Finally, the plasmid also played an important role in chlamydial adaptation to the large intestine. It took 10-fold more plasmid-deficient Chlamydia organisms than plasmid-positive Chlamydia organisms to achieve long-lasting colonization following intracolon inoculation. Furthermore, plasmid-positive Chlamydia improved colonization efficiency over time following intracolon inoculation, while plasmid-deficient Chlamydia failed to do so. Thus, the plasmid is able to improve chlamydial fitness in all regions of the GI tract but with the most significant role in the small intestine. Since different regions of the GI tract are equipped with different immune effectors (50–52) and the role of plasmid in improving chlamydial colonization in different regions varies considerably in terms of quantity and time kinetics, we propose that Chlamydia may have acquired the cryptic plasmid to improve its fitness in different regions of GI tract via different mechanisms. The fact that the plasmid both harbors 8 different genes and regulates two dozen chromosomal genes (56) suggests that the plasmid may have the capacity to do so. The present study has laid a foundation to further reveal the specific mechanisms by which different plasmid-dependent factors interact with different host effectors in different regions of the GI tract.
The finding that no infectious particles were ever recovered from small intestinal tissues of mice intragastrically inoculated with plasmid-deficient Chlamydia was a surprise to us. Nevertheless, plasmid-deficient chlamydial genomes were recovered from the same tissues, and the number of chlamydial genomes increased over time, suggesting that the plasmid-deficient chlamydial organisms were live but not infectious in the small intestine. This status is similar to that with persistent infection. C. muridarum can be induced to undergo persistence (57, 58). Both gastric and small intestinal effectors might contribute to the induction of plasmid-deficient Chlamydia into persistent infection in the small intestine, since a direct inoculation of plasmid-deficient Chlamydia into jejunum led to recovery of infectious particles. We have recently shown that the plasmid-encoded pGP3 plays an important role in promoting chlamydial survival in the stomach by resisting gastric acid (49). In the absence of plasmid, chlamydial organisms may use persistence as an alternative strategy for dealing with gastric acid stress. The chlamydial chromosome does carry acid stress response genes (59). It will be interesting to investigate how gastric and small intestinal effectors trigger and maintain chlamydial persistence. Regardless of how the noninfectious status was induced, this small intestinal infection condition may provide an ideal in vivo model for investigating the mechanism and evaluating the biological significance of chlamydial persistence.
A second surprising finding from the present study is that the cryptic plasmid is more important in promoting chlamydial colonization in the small intestine than in the large intestine. This conclusion is supported by the series of experiments that used quantitative comparison of plasmid-positive versus plasmid-deficient Chlamydia for colonizing small versus large intestinal tissues. Following intragastric or intrajejunal inoculation with plasmid-deficient Chlamydia, no or 530-fold fewer infectious organisms were recovered from the small intestine. However, when the same organisms were intracolonically inoculated, plasmid-deficient Chlamydia was only 7- to 120-fold less capable of establishing colonization. This difference may be caused by both different growth advantages of plasmid-positive versus -deficient Chlamydia and different effectors in small versus large intestines. Plasmid-deficient C. muridarum is known to be less efficient than plasmid-positive C. muridarum in colonizing mouse genital tract tissues, although both organisms can establish robust colonization (16). Thus, the reduced growth advantage by plasmid-deficient Chlamydia may contribute to its deficient colonization in different intestinal tissues. However, the different infectious particle recoveries of the same plasmid-deficient Chlamydia strain from small versus large intestines may be regulated by different intestinal effectors. For example, the small intestinal effectors might be able to drive chlamydial persistence, while large intestinal effectors failed to do so. It is worth noting that we have recently shown that C. muridarum can spread via the hematogenous route (41), which partially explains why when C. muridarum was introduced to mouse mucosae, it readily colonized the GI tract (37). Thus, chlamydial organisms inoculated into different regions of the GI tract may spread across the GI tract via a hematogenous route in addition to the lumenal spreading discussed above. It is worth differentiating the contribution of hematogenous spreading from that of lumenal spreading.
MATERIALS AND METHODS
Chlamydia muridarum organisms.Chlamydia muridarum strain Nigg3 was propagated and purified in HeLa cells (ATCC catalog number CCL2 [60]). Nigg3 genome sequence (GenBank accession CP009760.1) is different from that of the reference C. muridarum (26). Nigg3 was also used to derive a plasmid-deficient clone (designated CMUT3.G5 [13, 32]). Both Nigg3 (designated plasmid positive) and plasmid-deficient CMUT3.G5 were purified as elementary bodies (EBs) and stored in aliquots of SPG buffer (0.2 M sucrose, 20 mM sodium phosphate [pH 7.4], and 5 mM glutamic acid) at −80°C.
Mouse infection.All animal experiments were carried out in accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the Committee on the Ethics of Laboratory Animal Experiments of the University of Texas Health Science Center at San Antonio.
Six- to seven-week-old female C57BL/6J (stock number 000664; Jackson Laboratories, Inc., Bar Harbor, ME) mice were inoculated with purified C. muridarum EBs at different doses (measured in inclusion-forming units [IFU]) via different routes as indicated for individual experiments. After inoculation, mice were monitored for rectal shedding of live organisms or sacrificed for monitoring live organisms or chlamydial genomes in different tissues at designated time points after infection as indicated for individual experiments. The oral or intragastric inoculation was carried out by intragastric intubation of chlamydial organisms in a total volume of 200 μl SPG buffer using a straight ball-tipped needle (N-PK 020; Braintree Scientific, Inc., Braintree, MA) as described previously (48). Intrajejunal inoculation was carried out as described previously (61, 62). Briefly, mice anesthetized with isoflurane underwent laparotomy (0.5-cm incision). Chlamydial organisms in a total volume of 50 μl SPG buffer were injected into the middle region of jejunum lumen with a 27-gauge needle. The jejunum middle region was identified and pulled closer to the abdominal wall to aid the injection by using curved forceps. After gently pulling out the needle, the jejunum was returned to the abdominal cavity, and the abdominal wall was closed with two or three 9-mm autoclips using an autoclip applier (both from Braintree Scientific, Inc.). For intracolon inoculation, chlamydial organisms were diluted in 50 μl of SPG buffer containing the desired number of IFU as indicated for individual experiments and delivered to the colon using a straight ball-tipped needle designed for mouse oral gavage (N-PK 020; Braintree Scientific, Inc.). After inoculation, mice were monitored for rectal live organism shedding or sacrificed to titrate live organisms in corresponding samples.
Titrating live chlamydial organisms recovered from rectal swabs and tissues.To quantitate live chlamydial organisms in rectal swabs, each swab was vortexed with glass beads in 0.5 ml of SPG buffer. For the titrating of live chlamydial organisms recovered from mouse tissues, different GI tract tissues were harvested on designated days after inoculation as specified for individual experiments. Each tissue segment was transferred to a vial containing 2 ml cold SPG buffer including stomach (Sto), duodenum (Duo), jejunum (Jej), ileum (Ile), cecum (Cec), colon (Col), or anorectum (Rec) for homogenization and sonication (63). The chlamydial organisms released into the supernatants were titrated on HeLa cell monolayers in duplicates and counted using an immunofluorescence assay (64). The total number of IFU per swab was calculated based on the mean IFU per view, the ratio of the view area to that of the well, dilution factor, and inoculation volumes. Where possible, a mean IFU/swab was derived from the serially diluted samples for any given swab. The total numbers of IFU/swab were converted into log10 for calculating group means and standard deviations.
Titrating the number of C. muridarum genomes in mouse tissues using quantitative PCR.To quantitate the genome copies of C. muridarum in a given tissue sample, 200 μl of each tissue homogenate prepared as described above (prior to sonication) was transferred to a lysis buffer provided with the Quick-gDNA miniprep kit (catalog number 11-317C; Genesee Scientific, San Diego, CA) for DNA extraction according to the manufacturer’s instructions. Each DNA preparation was eluted in 100 μl elution buffer, and 2 μl was used for quantitative PCR (qPCR). The following primers derived from the chlamydial 16S rRNA coding region were used: forward primer (5′-CGCCTGAGGAGTACACTCGCAGGA-3), reverse primer (5′-CCAACACCTCACGGCACGAG-3′), and double-quenched probe (5′-CACAAGCAGTGGAGCATGTGGTTTAA-3′) (Integrated DNA Technologies, Coralville, IA). PCRs were carried out in a total volume of 10 μl in a CFX96 Touch deep-well real-time PCR Detection system with IQ Supermix real-time PCR reagent (Bio-Rad, Hercules, CA). The qPCR conditions included an initial denaturation step at 95°C for 3 min, followed by 40 cycles of amplification at 95°C for 15 s and 60°C for 1 min. Genome copy numbers from a given sample in triplicates were calculated based on a standard plasmid DNA and expressed as log10 genomes per sample.
Statistics analyses.The numbers of live organisms (in IFU) and genome copies either at individual data points or over a time course were compared using Wilcoxon rank sum test. Areas under the curves (AUCs) were used for comparing time course data. The 50% infection dose (ID50) was calculated using the IC50 calculator from AAT Bioquest (https://www.aatbio.com/tools/ic50-calculator/) based on IFU positivity in mouse samples (49).
ACKNOWLEDGMENTS
This work was supported in part by grants (to G. Zhong) from the U.S. National Institutes of Health. J.M. is supported by the Tianjin Medical University General Hospital.
FOOTNOTES
- Received 9 November 2019.
- Returned for modification 25 November 2019.
- Accepted 20 December 2019.
- Accepted manuscript posted online 23 December 2019.
- Copyright © 2020 American Society for Microbiology.
