SUBVERSION OF THE EUKARYOTIC HOST CELL BY SECRETED CHLAMYDIAL EFFECTOR PROTEINS
Ted Hackstadt
Host-Parasite Interactions Section, Laboratory of Intracellular Parasites, National Institutes of Health, National Institute of Allergy and Infectious Diseases, Hamilton, MT, USA
Chlamydiae occupy a unique vacuolar niche within the host cell. The chlamydial inclusion, unlike vacuoles containing other intracellular pathogens, is not interactive with endocytic vesicular trafficking pathways but is instead fusogenic with an incompletely understood exocytic pathway which otherwise delivers sphingomyelin and cholesterol from the Golgi apparatus to the plasma membrane.
Entry into this pathway is an active process on the part of the chlamydiae as both de novo transcription and translation are required. As obligate intracellular pathogens intimately dependent upon the eukaryotic host cell, chlamydiae clearly communicate with the host cell in many ways to promote pathogen survival and replication. Many Gram-negative pathogens modify cellular functions through a targeted type m secretion apparatus that delivers bacterial effector proteins directly into the cytoplasm of the host cell [1], Analysis of the chlamydial genome has revealed the capacity to encode a complete type m secretion system [2-5]. Proteomic analyses have indicated a functional type EI secretion apparatus on EBs [6]. Secretion of type IH effector proteins are now recognized as potential controlling elements for many significant events during the chlamydial developmental cycle. Some of the major questions remaining in chlamydial biology are dependent upon when the secretion apparatus is active and the functions of the secreted effectors.
Many of the changes in the interactions of the nascent inclusion require chlamydial protein synthesis and are specific and localized to the inclusion. This specificity strongly suggests modification of the exposed inclusion membrane. Examples of chlamydial proteins controlling localized events could include the
recruitment of actin to promote entry [7-10].
Other examples of cis-acting modifications to the nascent inclusion membrane may include:
evasion of lysosomal fusion [11-15], interactions with microtubules to deliver the nascent inclusion to the peri-Golgi region and microtubule organizing center [16-18], initiation of fusion with exocytic vesicular traffic from the Golgi apparatus [14, 19-21], and recruitment of, but not fusion with, recycling endosomes containing transferrin and its receptor [15, 22-24], Examples-of-more trans-acting chlamydial control of host cell functions include inhibition of apoptosis [25-29], activation of caspase-1 [30], down-regulation of MHC class I and II expression
[31, 32], activation of the Raf/MEK/ERK/cPLA2 signaling pathway [33], and alterations of host transcriptional profiles [34, 35],
Attachment of chlamydiae to host cells and initiation of the entry process has been one of the more controversial and interesting areas of chlamydial pathogenesis. A large number of chlamydial ligands and host receptors have been proposed to play a role in this essential process [36]. Recent studies with chemically mutagenized CHO cells have distinguished two distinct stages in the chlamydial entry process [37, 38] Heparin sulfate like proteoglycans are believed to play a role in an initial, electrostatic and reversible interaction that precedes an irreversible step leading to internalization [39, 40]. The host gene mutated to block the secondary, irreversible step has not been identified. The host responses and requirements for chlamydial entry are, however, beginning to be described. One of the first overt responses of the host cell is the recruitment of actin to the site of EB attachment [8]. This results in the formation of
a pedestal-like structure with the EB at the apex. This recruitment of actin occurs rapidly and culminates in the internalization of the EB.
Host signaling molecules known to be recruited and required include the RAC GTPase [9], WAVE2, and the Arp2/3 complex. A chlamydial type EH secreted effector has been proposed to play an important role in the initiation of this process [10]. The protein, named Tarp for Translocated Actin Recruiting Phosphoprotein, is pre-existing in an unphosphorylated form in EBs and tyrosine phosphorylated within minutes of attachment to susceptible host cells. Although the molecular mechanisms triggering cell signaling pathways remain to be defined, Tarp appears to be translocated to the cytoplasmic face of the plasma membrane and tyrosine phosphorylated while the EBs are still extracellular. Whether this protein actually serves as a receptor aiding chlamydial attachment to the host cell or is simply a signaling molecule remains to be determined.
The entire process of cell signaling, actin recruitment, and internalization of EBs is transient and over within minutes [9].
Endocytosed EBs are then found within a plasma membrane derived vesicle that is effectively dissociated from all cellular processes and displays no known cellular markers [15]. This vacuole remains non-interactive until chlamydial protein synthesis is initiated. Once chlamydial protein synthesis has begun, the interactions with the host cell change dramatically.
While the chlamydial inclusion has been known to be non-lysosomal in character for some time now [41], the properties of the vacuole and relative contributions of the host and parasite to its establishment are only recently beginning to be deciphered. Surveys of early and late endosomal markers, lysosomal markers, fluid phase, and plasma membrane markers demonstrate no interaction of the endocytic pathway with the inclusion membrane of viable chlamydiae [13, 42], Instead, the chlamydial inclusion acquires
sphingomyelin and cholesterol directly from the Golgi apparatus from what appears to be an exocytic pathway which normally delivers these lipids to the plasma membrane. Delivery of sphingomyelin and cholesterol is time, temperature, and energy dependent and is blocked by inhibitors of anterograde Golgi trafficking, such as brefeldin A, or of microtubule function, such as nocadazole [19-21]. Recognition of this pathway has provided many insights into the cellular interactions of the chlamydial inclusion.
All species of chlamydia interact similarly with this pathway to acquire sphingomyelin [19, 43, 44]. The molecular mechanisms employed by chlamydiae to subvert cellular trafficking pathways to the advantage of the parasite remain largely unknown although they almost certainly involve specific modification of the inclusion membrane. Most Golgi functions, such as glycosylation and transport of proteins, are not measurably disrupted in C. trachomatis-infected cells [22], Indeed, even in chlamydia-infected cells co-chlamydia-infected with Coxiella burnetii, sphingomyelin is delivered only to the chlamydial inclusion and not to the C.
burnetii parasitophorous vacuole [13]. The chlamydial inclusion is therefore specifically recognized as a target by the cellular machinery controlling sphingomyelin trafficking. Initiation of fusion with sphingomyelin-containing vesicles is initiated within about 2 hr following internalization of C. trachomatis. In the presence of inhibitors of bacterial transcription or translation, endocytosed EBs do not attain the ability to acquire sphingomyelin and remain within vesicles that are effectively non-fusogenic with any cellular organelles [14], The collective interpretation of these observations is that chlamydiae display two distinct mechanisms for evasion of lysosomal fusion. One, which until recently had been considered passive or mediated by structural components of EB, does not require chlamydial protein synthesis and has been proposed to confer a survival advantage upon EBs until such time as they
initiate protein synthesis. A more active evasion of lysosomal fusion involves chlamydial modification of the inclusion membrane and is characterized by the ability to incorporate sphingomyelin. It has been proposed that once the properties of the inclusion have been established, the chlamydiae are essentially sequestered in a vesicle whose fusion properties are incompatible with the endosomal/lysosomal pathway thus providing a favorable intracellular site for their replication.
The molecular mechanisms underlying chlamydial redirection of vesicular fusion remain unclear although the requirement for chlamydial protein synthesis and the specific alteration of only the inclusion membrane suggests that chlamydiae modify the cytoplasmic face of the inclusion, likely by the insertion of chlamydial protein. Concomitant with the change in fusogenicity is a microtubule-dependent trafficking of the nascent inclusion to a perinuclear region in close proximity to the Golgi apparatus [16-18].
Also during this early time frame, nascent inclusions begin to recruit early endosomes recycling transferrin and its receptor into very close apposition with the inclusion membrane although there is no fusion with these vesicles [14, 15, 23, 24]. Each of these processes require chlamydial protein synthesis and specific modification of the cytoplasmic face of the inclusion membrane although none of the chlamydial proteins controlling these events have been identified and it is not clear whether they are mediated by the same or different proteins.
An entire class of chlamydial proteins modifying the inclusion membrane is now recognized. It is clear that chlamydiae modify the inclusion membrane extensively with a large number of inclusion membrane proteins, collectively referred to as Incs . At least forty inclusion membrane proteins have been described in C. trachomatis with about twenty confirmed by immunofluorescence with specific antibodies. The Inc protein family
shares little primary sequence similarity among members but is characterized by a long, 40 amino acids or more, predicted hydrophobic domain [45-49]. Exposure of a number of Incs on the cytoplasmic face of the inclusion has been demonstrated by microinjection of specific antibodies although the function(s) of the Inc proteins are largely unknown [50]. The majority of Incs are expressed early in the developmental cycle which is consistent with possible roles as mediators of vesicle fusion.
All Inc proteins thus far tested in heterologous secretion systems are secreted via type IH secretion [6, 51]. One would suspect that chlamydial proteins controlling interactions with the host cell would be highly similar, however, there is surprisingly little conservation of Incs between chlamydial species despite all interacting with the same host vesicular trafficking pathways. Either there are unrecognized conserved small domains that control these interactions or other, possibly more peripherally associated, proteins function in there capacities.
Chlamydial Tarp and the Inc proteins identify at least two distinct stages in chlamydial development where secreted effectors may play important roles in defining the outcome of infection. Certainly there are many other levels of communication between chlamydiae and the eukaryotic host cell.
Identifying the remaining secreted effector molecules and their functions may be challenging but will continue to provide insights into the many adaptations chlamydiae utilize as successful parasites.
References
1. Hueck, C.J., Microbiol Mol Biol Rev, 1998. 62:379.
2. Hsia, R.C., et al., Mol Microbiol, 1997.
25:351.
3. Stephens, R.S., et al., Science, 1998.
282:754.
4. Read, T.D., et al., Nucleic Acids Res, 2000. 28:1397.
5. Subtil, A., A. Blocker, and A. Dautry-Varsat, Microbes Infect, 2000. 2:367.
6. Fields, K.A., et al., Mol Microbiol, 2003.
48:671.
7. Ward, M.E. and A. Murray, J Gen Microbiol, 1984. 130:1765.
8. Carabeo, R.A., et al., Infect Immun, 2002.
70:3793.
9. Carabeo, R.A., et al., Traffic, 2004. 5:418.
10. Clifton, D.R., et al., Proc Natl Acad Sci U S A, 2004. 101:10166
11. Wyrick, P.B., E.A. Brownridge, and B.E.
Ivins, Infect Immun, 1978. 19:1061.
12. Eissenberg, L.G. and P.B. Wyrick, Infect Immun, 1981. 32:889.
13. Heinzen, R.A., et al., Infect Immun, 1996.
64:796.
14. Scidmore, M.A., et al., Infect Immun, 1996. 64:5366.
15. Scidmore, M.A., E.R. Fischer, and T.
Hackstadt, Infect Immun, 2003. 71:973.
16. Higashi, N., Exp. Mol. Pathol., 1955.
4:24.
17. Clausen, J.D., et al., Mol Microbiol, 1997.
25:441.
18. Grieshaber, S., N. Grieshaber, and T.
Hackstadt, J Cell Biol, 2003. 116:3793.
19. Hackstadt, T., M.A. Scidmore, and D.D.
Rockey, Proc Natl Acad Sci USA, 1995.
92:4877.
20. Hackstadt, T., et al., EMBO J, 1996.
15:964.
21. Carabeo, R.A., D.J. Mead, and T.
Hackstadt, Proc Natl Acad Sci USA, 2003. 100:6771.
22. Scidmore, M.A., E.R. Fischer, and T.
Hackstadt, J Cell Biol, 1996. 134:363.
23. van Ooij, C., G. Apodaca, and J. Engel, Infect Immun, 1997. 65:758.
24. Raulston, J.E., Infect Immun, 1997.
65:4539.
25. Fan, T., et al., J Exp Med, 1998. 187:487.
26. Geng, Y., et al., J Immunol, 2000.
164:5522.
27. Fischer, S.F., et al., Infect Immun, 2001.
69:7121.
28. Rajalingam, K., et al., Infect Immun, 2001.
69:7880.
29. Schoier, J., et al., Microb Pathog, 2001.
31:173.
30. Lu, H„ C. Shen, and R.C. Brunham, J Immunol, 2000. 165:1463.
31. Zhong, G., T. Fan, and L. Liu, J Exp Med, 1999. 189:1931.
32. Zhong, G., et al., J Exp Med, 2000.
191:1525.
33. Su, H., et al., J Biol Chem, 2004.
279:9409.
34. Coombes, B.K. and J.B. Mahoney, Infect Immun, 2001. 69:1420.
35. Xia, M., et al., J Infect Dis, 2003. 187:424.
36. Hackstadt, T., Cell Biology., in Chlamydia: Intracellular biology, pathogenesis, and immunity., R.S.
Stephens, Editor. 1999, ASM Press:
Washington, D. C. p. 101.
37. Carabeo, R.A. and T. Hackstadt, Infect Immun, 2001. 69:5899.
38. Fudyk, T„ L. Olinger, and R.S. Stephens, Infect Immun, 2002. 70:6444.
39. Zhang, J.P. and R.S. Stephens, Cell, 1992.
69:861.
40. Su, H., et al., Proc Natl Acad Sci USA, 1996. 93:11143.
41. Wyrick, P.B. and E.A. Brownridge, Infect Immun, 1978. 19:1054.
42. Taraska, T., et al., Infect Immun, 1996.
64:3713.
43. Rockey, D.D., E.R. Fischer, and T.
Hackstadt, Infect Immun, 1996. 64:4269.
44. Wolf, K. and T. Hackstadt, Cell Microbiol, 2001. 3:145.
45. Rockey, D.D., R.A. Heinzen, and T.
Hackstadt, Mol Microbiol, 1995. 15:617.
46. Bannantine, J.P., D.D. Rockey, and T.
Hackstadt, Mol Microbiol, 1998. 28:1017.
47. Scidmore-Carlson, M.A., et al., Mol Microbiol, 1999. 33:753.
48. Bannantine, J.P., et al., Cell Microbiol, 2000. 2:35.
49. Shaw, E.I., et al., Mol Microbiol, 2000.
37:913.
50. Hackstadt, T., et al., Cell Microbiol, 1999.
1:119.
51. Subtil, A., C. Parsot, and A. Dautry-Varsat, Mol Microbiol, 2001. 39:792.
THE GENERATION TIME OF CHLAMYDIA TRACHOMATIS
I. Miyairi1'2*, O.S. Mahdi3, S.P.Ouellette3, R. Beiland3, G.I. Byrne3
Departments of 'infectious Diseases, St. Jude Children's Research Hospital, Lauderdale, Memphis TN and department of Pediatrics,3 Molecular Sciences, U. Tennessee Health Science Center, Memphis, TN USA
The generation time of chlamydia has previously been estimated from results of quantitative gene expression assays1'2, however this may not represent the rate of actual cell division. We sought to determine the generation time of Chlamydia trachomatis by arresting cell division sequentially with addition of penicillin and enumerating aberrant RBs within an inclusion under light microscopy3. A one step growth curve for elemental bodies was created for comparison.
HeLa 229 epithelial cells were grown on 8 well chamber/slides and infected with C.
trachomatis serovar D (MOI 0.3). 50 microgram/ml of penicillin was added to quadruplicate samples at 0, 6, 12, 15, 18, 19.5, 21, 24, or 30 hours post infection (hpi). SPG was added as a control at 0 hpi. C. trachomatis and infected cells were fixed with methanol, stained with Giemsa fluid at 42 hpi and examined by oil immersion light microscopy.
The number of large aberrant RBs in each inclusion was counted and averaged for 200 inclusions per sample. A one step growth curve was also produced by determining the number of inclusion forming units in replicate samples. HeLa 229 cells were grown on 24 well plates, infected with C. trachomatis serovar D (MOI 0.3) in triplicates and collected at 6,12, 15, 18, 21, 24, 30, 39, 42 hpi.
Cell suspensions were homogenized for IFU assay.
The average number of RBs (standard deviation) at 6, 12, 15, 18, 19.5 hpi was 1.1(0.3), 1.5(1.2), 3.8(3.8), 8.1(6.2), 14.0(10.0), respectively. We were unable to -enumerate beyond 21 hpi because of overcrowding. The lag phase was observed to
be approximately 12 hours and the generation time was 2.4 hours, for the initial four rounds of replication. No infectivity was demonstrable for the first 21 hpi, and at 24, 30, 39, 42 hpi was 1.0(0.3) xl0*5, 1.6(0.3) xl0*6, 1.4(0.2) xl0*7, and 1.5(0.2) xl0*7 respectively. 16S rRNA transcript levels also will be measured to create a corresponding genomic generation curve for comparison.
The generation time of C. trachomatis averaged 2.4 hours corresponding well to previous estimates by quantitative gene expression assays1'2. There was a large distribution of the number of RB per inclusion, suggesting replication occurs asynchronously.
The lag phase was estimated to be about 12 hours but may be up to 18 hours in significant population of organisms. This extended lag phase has not been previously noted.
References
1. Alexander JJ. Effect of infection with the
meningopneumonitis agent on deoxyribonucleic acid and protein synthesis by
its L-cell host. J Bact. 97; 653-57.
2. Wilson DP, Mathews S, Wan C, Pettitt AN, McElwain DL. Use of a quantitative gene expression assay based on micro-array techniques and a mathematical model for the investigation of chlamydial generation time.
Bull Math Biol. 2004 May; 66(3):523-37.
3. Matsumoto A, Manire GP. Electron microscopic observations on the effects of penicillin on the morphology of Chlamydia psittaci. J Bact. 101; 278-285.
A TYPE THREE SECRETION/CONTACT-DEPENDENT MODEL OF CHLAMYDIAL INTRACELLULAR DEVELOPMENT
D.P. Wilson1, P. timms2, D.L.S. Mcelwain3 and P.M. Bavoil4*
1 2 Department of Biomathematics, University of California, Los Angeles, California, USA; School
of Life Sciences & 3School of Mathematical Sciences, Queensland University of Technology, Brisbane, Queensland, Australia; 4Department of Biomedical Sciences, University of Maryland Dental School, Baltimore, USA
Late in development, Chlamydia's replicative form, the reticulate body (RB), differentiates into the infectious form, the elementary body (EB). The signal(s) that trigger this crucial physiological change are unknown.
We present a hypothesis for the modulation of chlamydial late developmental events involving the contact-dependent type HI secretion (TTS) system. In this model, TTS surface projections mediate intimate contact between the RB and the inclusion membrane.
Below a threshold number of projections, detachment of the RB provides a signal for late differentiation of RB into EB.
We use published data and our own observations to develop a mathematical model investigating this hypothesis. If the hypothesis proves to be accurate, then increasing the number of inclusions per host cell will increase the number of infectious progeny EB.
Conversely, TTS down-regulation resulting in a reduction of the number of projections on the surface of the RB or TTS selective inactivation will significantly reduce the burst size of infectious chlamydial particles. The full implications of the hypothesis as predicted by the model provide a ground for experimental verification.
DECIPHERING CHLAMYDIAL GENE REGULATION BY IN SILICO PREDICTION OF PROMOTER SEQUENCES
Peter Timms1, Brian Grech1 and Sarah A Mathews1
'School of Life Sciences, Queensland University of Technology, Brisbane, Australia Even though Chlamydia has a relatively
small genome, remarkably little is known about gene regulation in this important human and animal pathogen. The three chlamydial sigma factors, sigma 66, sigma 28 and the alternate sigma factor, sigma 54 are differentially expressed during the chlamydial developmental cycle, suggesting an involvement in stage-specific regulation of the chlamydial transcriptome. Prior to 1998, only 22 chlamydial promoters had been studied at the level of transcript start site mapping or by using in vitro transcription assays. With the full genome sequence of over eight chlamydial strains now available, bioinformatic approaches can be utilized effectively to predict chlamydial promoters. We have applied advanced bioinformatics approaches to searching the whole chlamydial genomes to predict promoters.
We developed an in silico whole genome analysis approach to search for promoters in the chalmydial genome. This involved the construction of a percentile weighted matrix to represent the -35 and -10 promoter hexamers for both C.
trachomatis and C. pneumoniae. We searched the whole chlamydial genomes using these weighted matrices and selected motif hits with scores above a threshold. We then reduced the number of predictions by phylogenetic footprinting between C. trachomatis and C. pneumoniae. We confirmed our in silico outputs by mapping the transcript start sites of selected genes.
When the whole C. trachomatis genome was screened with the sigma 70 promoter weighted matrix (based on E. coli sigma 70 data), 1000's of potential sigma 66-type promoters were predicted, many of which were clearly false positives.
Phylogenetic footprinting of the in silico predicted promoters between the available chlamydial genome sequences reduced the number to 36 highly conserved promoters, sigma 66-type chlamydial promoters. We then analysed 14 of these promoters via transcript start site mapping of the relevant gene and were able to confirm that our in silico prediction was correct in 7% of cases.
This in silico approach is slightly biased, as it uses the E.coli sigma 70 data as the basis of the matrix.
Nevertheless, the consensus in silico chlamydial sigma 66 promoter TTGATT TATAAT (bold indicates high base preference) does show some variations when compared to the E. coli sigma 70 consensus of TTGACA TATAAT. Interestingly, the chlamydial in silico consensus also varies compared to the consensus produced by the 36 currently mapped chlamydial promoters TTGAXX TATAAT.
In silico prediction of promoters in whole chlamydial genomes can be a useful means to predict new promoters and should play an important role in unraveling our understanding of the subtle and potentially complex gene regulatory networks that enable this unique pathogen to be so successful. Our in silico approach was strongly biased towards the E. coli sigma 70 type promoter consensus, however it did predict slight variations in the chalmydial genomes analysed. Our prediction success rate of over 70% is extremely good for predicting further novel promoters in Chlamydia.
Interestingly, a significant number of biologically confirmed promoters in one chlamydial species did not readily footprint across into the other chlamydial species. It is unknown why so many promoters are different between the chlamydial species, but this may suggest significantly different mechanisms of regulating key genes between the species. Other interesting features emerging from chlamydial genome analysis are that several genes appear to have multiple promoters, often from two different sigma factor classes and some genes have promoters located within the coding region of the upstream gene. This suggests that multiple levels of gene regulation exist for some different classes of genes. Overall, in silico prediction of promoter hexamers and other transcription factor binding sites should play an important role in defining the mechanisms of gene regulation used by this unique intracellular parasite.
THE (^-GLUTAMINE S-ADENOSYL-L-METHIONINE DEPENDENT METHYL-TRANSFERASE PRMC/HEMK IN CHLAMYDIA TRACHOMATIS METHYLATES CLASS 1 RELEASE FACTORS
Yvonne Pannekoek1*, Valerie Heurgué-Hamard2, Ankie A.J. Langerak1 Dave Spijer3, Richard Buckingham and Arie van der Ende1.
Academic Medical Center, department of Medical Microbiology and department of Biochemistry, University of Amsterdam, Amsterdam, The Netherlands and 2Institut de Biologie Physico-Chimique, Service de Biochimie, CNRS, Paris, France
Methylation of DNA and protein plays an important role in gene expression and activity of enzymes. Methylation is catalyzed by methyltransferases (MTases) using S-adenosyl- L-methionine (AdoMet) as methyl donor. The role of methylation in the regulation of gene expression and protein activity of the obligate intracellular pathogen Chlamydia trachomatis is unknown.
Genomes of Chlamydia spp. apparently lack the homologues of genes encoding well characterized DNA MTases, as well as the gene enabling the synthesis of AdoMet. This raises the question whether methylation events in Chlamydia spp. do occur. Recently, PrmC has been demonstrated to act in E. coli as an /V5-Glutamine AdoMet dependent MTase of class I peptide release factors (RFs), thereby making PrmC the first N5 -Glutamine MTase identified (Heurgue-Hamard et al EMBO J 2002 (4): 769-78). We used a genetic approach to identify and analyze the putative prmC homologue of C.
trachomatis.
CT024 (prmQ and chlamydial RFs (prfA and prfB) of C. trachomatis strain L2 were PCR amplified and cloned into the expression vector pQEL-80 thereby creating His-tagged fusion proteins. Next, the genes of prfA and prfB as well as the DNA encoding
the His tag were subcloned into the low copy number pWSK129. pWSK129 is compatible with pQEL-80. Methylation of RFs was analyzed by MALDI-TOF mass spectrometry.
Neither RF1 nor RF2 is methylated in E. coli prmC knock outs, leading to global deficiency in termination of translation and hence a deliberated growth. Overexpression of K12 RF2, thereby overwhelming PrmC, also leads to impaired growth. Taking advantage of these phenotypes, potential PrmC function of CT024 was assessed in the genetic background of E. coli.
Expression of CT024 in a E. coli prmC knock out demonstrated that CT024 restores the growth defect of this strain, suggesting an interaction of CT024 with RFs of E. coli.
This was substantiated by the observation that overexpression of CT024 suppressed the toxic effect of overexpression of K12 E. coli RF2 and strongly suggested that CT024 methylates RFs. Indeed, in vivo methylation assays carried out with recombinant CT024 and RFs of chlamydial origin demonstrated that CT024 methylates RFs within the tryptic fragment containing the universally conserved domain Gly-Gly-Gln. This is Completely consistent with the enzymatic properties of PrmC of E. coli origin.
We conclude that CT024 encodes a functional chlamydial PrmC, acting as an N5 -Glutamine AdoMet dependent MTase of RFs.
This makes PrmC the first MTase of C.
trachomatis demonstrated to be involved in the basic biological process of translation termination.
Studies are now underway to identify the putative chlamydial AdoMet permease.