Supporting information for Blocker et al. (2003) Proc. Natl. Acad. Sci. USA, 10.1073/pnas.0535335100

 

Supporting Text

Structure

Old and New Sequence Homologies. We find that functional homologs in TTSS and flagella can often be recognized in advance by matching structural characters (size, pI, and "instability index," found in SWISSPROT, ref. 1), with prior knowledge about the proteins, such as whether and when they are secreted in the assembly pathways. A recently understood example is InvJ (Salmonella SPI1, ref. 2)/Spa32 (Shigella, refs. 3 and 4), which works in a manner similar to flagellar FliK (5, 6) to regulate the transition from the needle/hook to effector/flagellar filament component secretion.

Morphological Divergence. The TTS machines of Salmonella and Shigella are dispersed over the entire bacterial surface (50–100 copies per cell, ref. 7) whereas the Hrp pili of Ralstonia solanacearum emanate from only one pole of the bacterium, presumably the pole at which they also attach to the plant cell (8). Some TTS machines need to traverse long distances before reaching their host cell membrane. Enteropathogenic Escherichia coli (EPEC) attaches to the apical face of enterocytes. Its TTSS must traverse the mucus, glycocalyx and disrupt the microvilli to reach the apical plasma membrane of the host cell. EPECs have a long TTSS-encoded protein conduit, the EspA filament synthesized near the host cell, which connects bacterium and cell (9, 10). EPEC also possess an NC, the needle of which is composed of the PrgI/MxiH homolog EscF (11), connected at its distal tip to the wider EspA filament (12, 13). Salmonella have other surface appendages called "invasomes" (14) but their relation to NCs remains unclear because their dependency on SPI1 is controversial (15). Perhaps Salmonella needs these extra appendages to invade epithelial cells (with microvilli) in addition to M cells (no microvilli and active luminal transcytosis)? The TTS machines of plant pathogens must traverse the host cell wall, which they do by using the Hrp pilus (8, 16). There is a potential needle protein equivalent in plant pathogens (1). Thus, some TTS machineries polymerize an additional structure on top of the needle, much like the flagellar filament elongates from the hook, upon encountering signals indicating host cell vicinity and may only secrete effectors once tight contact with the host cell has been established.

Regulation

Regulation of Expression. In Salmonella SP1, HilD and HilC (class I-like?) act on HilA, which acts on InvF and the majority of genes encoding the NC (17–20). The HilA box is found at these promoters (class II-like?, ref. 21). In the plant pathogens Pseudomonas and Erwinia, an hrp box (22–24), a cis-acting DNA element that is recognized by the HrpL sigma factor (class II-like?, refs. 25 and 26) is found ≈50 bp upstream of the initiation codon of both TTSS- and putative effector-encoding operons. Those Xanthomonas spp. and Ralstonia solanacearum hrp genes that are regulated by the AraC-like regulator HrpX/HrpB contain a PIP box (plant inducible promoter; class II-like?, ref. 27). HrpX/B seems topped by HrpG, an OmpR-like two-component response regulator, acting perhaps as the basal master switch (class I-like?, ref. 28).

Chaperones as a Bridge Between Posttranslational and Cotranslational Secretion: Evidence for Regulatory Chaperones from Virulence TTSSs. Shigella IpgC is equivalent to SicA of Salmonella SPI1 (binds SipC and SipB) and LcrH/SycD of Yersinia (binds YopD). SicA can bind InvF, a transcriptional regulator of the SPI1 genes and the complex activates transcription of effector-encoding genes (29, 30). A sicA null mutation abolishes transcription of sopE (encoding a SPI1 effector; ref. 31). There are also posttranslational and cotranslational pathways for Yersinia effector Yop secretion (32). YopE is synthesized at 37ºC, has SycE as chaperone, and can apparently use both routes (but uses mostly the posttranslational one initially, ref. 33), whereas YopQ/K is only synthesized at 37ºC upon activation of secretion, has no known chaperone, and can only be secreted cotranslationally. LcrH and YopD have negative effects on late effector yop expression. The LcrH/YopD interaction is required for this but LcrH has a regulatory role in addition, which may involve its interaction with another chaperone-like protein, YscY (34). The LcrH/YopD complex binds the yopQ mRNA and is involved in its degradation under conditions when its expression is repressed (35). In Yersinia, a negative transcriptional regulator is known, LcrQ/YscM, but it is not thought to be an anti-σ factor nor conserved in other TTSSs.

Function

The ATPase: Energy Requirements and Transport Intermediates. Are effector motifs recognized by the secreton? The structure of the first 129 aa of YopH has been solved at atomic resolution, demonstrating the presence of the N-terminal amphipatic α-helix (36-38). One crystal structure is a dimer with swapped domains at the N termini, perhaps mimicking the SycH/YopH interaction and "preopening" YopH for export (37). The other is a monomer where the N terminus is folded back toward the rest of the protein (36), as in the NMR structure (38). This suggests that the N terminus of chaperoned effectors adopts an open or a closed position depending on the binding of the chaperone. Indeed, the structure of chaperone binding region of the Salmonella effector SptP bound to its chaperone SicP reveals that amino acids 35–139 of SptP are wound around two chaperone molecules (see also dimeric SycE structure, ref. 39) and thus maintained in an extended conformation (40). Yet, the remainder of effectors is probably folded when chaperones-bound because YopE shows its GAP activity when bound to SycE (41).

How is energy transduced by the export motor during secretion? Only the F1 hexamer has strong ATPase activity, monomers and dimers do not. Catalytic cooperativity is due to rotation of the asymmetric γ-subunit, driven by proton translocation in the F0 portion, sequentially causing ADP+Pi binding, interconversion to and release of ATP (42, 43). F1 is nearly 100% efficient (44). Accordingly, it synthesizes ATP in the presence of a proton gradient and hydrolyses it in its absence. Would interactions of the ATPase with substrates occur with protein subdomains or regions of extended carbon backbone at regular intervals? Might the correct export direction be dictated by the nucleotide bound in each subunit? Would the substrate crawl up the secreton channel at each ATP hydrolyzed? Currently, there is no evidence in support of our model. Yet, the structure of the ATPase TrwB involved in bacterial conjugation displays an identical homohexameric fold, despite lack of sequence homology to F1-type ATPases (45).

Translocation. Evidence for a pore. Pore protein insertion is kinetically, and thus probably energetically, coupled to their secretion from bacteria in the presence of biological membranes for Shigella, but not for EPEC (46, 47). Most cells types are injected by the Yersinia Ysc, but with varying efficiency. Knockout of the surface adhesins Invasin or YadA, abrogates type III injection into some, but not all cultured cells (48). For Salmonella and Shigella, there is no evidence for a pore receptor, whereas EPEC injects by using its TTSS, its own receptor, Tir, which binds its intimin adhesin (49). Purified Salmonella SipB (YopB/IpaB homolog) assembles into hexamers via an N-terminal domain forming a trimeric coiled-coil (50). It inserts into artificial lipid vesicles with its N and C termini on the presumed extracellular side, a large loop in the vesicle lumen and two transmembrane helices tilted in the membrane (51). A single pore-like structure putatively consisting of EspB and EspD has been visualized by atomic force microscopy (47). It is very large (>50 nm in outer diameter) and contains a central opening of 8 nm, although this may not be its minimal inner diameter. It is formed of six to eight repeating moieties. Although it is not clear from which side of the membrane the complex was viewed, it appears of asymmetric height (15–20 nm) on that side.

Modifications of the pore. Pore size regulation may be important where the TTSS-host cell interaction is long-lived (i.e., Yersinia Ysc, EPEC, and plant pathogens TTSSs all used to multiply at the surface of cells, and Salmonella SPI2 and Chlamydia TTSSs required for intravacuolar growth) rather than rapid (e.g., SPI1 in Salmonella and Shigella TTSS-mediated cell invasion). For the Yersinia Ysc, there is evidence that at least two TTSS-secreted proteins are involved, YopK and LcrV (52, 53). YopK is essential for virulence (53). A yopK mutant strain displays a higher cytotoxicity and contact hemolytic activity and a larger apparent pore size, whereas a yopK overexpressing strain displays the inverse (54). YopK localizes at the site of bacterium-host contact. LcrV is present at the bacterial surface before host cell contact and essential for virulence and Yop translocation. Antibodies against LcrV block Yop translocation (or pore insertion? ref. 55). Purified LcrV (which has no obvious hydrophobic regions) was shown to induce channels of 3 nS conductance in lipid bilayers. Does LcrV associate with the YopB/YopD in the host membrane and regulate pore opening?

Finally, how the pore might be disassembled by organisms, like Shigella, which release themselves into the host cytoplasm or cause oncosis (Pseudomonas, refs. 56 and 57) remains to be addressed.

Continuity among needle, extension filament, and pore. Morphological evidence for continuity between the needle and the extension filament of those organisms that carry them was provided for EPEC (12, 13). There is also evidence for continuity between the EspA filament and the EspB pore component, shown to interact directly by several means (58). Binding of EspA filaments to host cells occurs normally in the absence of EspB, suggesting that after initial attachment of the EspA filaments EspB is delivered into the host cell membrane and that the EspA-EspB interaction is important during effector translocation.

1. Aizawa, S.-I. (2001) FEMS Microbiol. Lett. 202, 157–164.

2. Kubori, T., Sukhan, A., Aizawa, S.-I. & Galan, J. E. (2000) Proc. Natl. Acad. Sci. USA 97, 10225–10230.

3. Tamano, K., Katayama, E., Toyotome, T. & Sasakawa, C. (2002) J. Bacteriol. 184, 1244–1252.

4. Magdalena, J., Hachani, A., Chamekh, M., Jouihri, N., Gounon, P., Blocker, A. & Allaoui, A. (2002) ) J. Bacteriol. 184, 3433–3441.

5. Williams, A. W., Yamaguchi, S., Togashi, F., Aizawa, S. I., Kawagishi, I. & Macnab, R. M. (1996) J. Bacteriol. 178, 2960–2970.

6. Muramoto, K., Makishima, S., Aizawa, S.-I. & Macnab, R.-M. (1998) J. Mol. Biol. 277, 871–882.

7. Blocker, A., Gounon, P., Larquet, E., Niebuhr, K., Cabiaux, V., Parsot, C. & Sansonetti, P. (1999) J. Cell Biol. 147, 683–693.

8. Van Gijsegem, F., Vasse, F., Camus, J.-C., Marenda, M. & Boucher, C. (2000) Mol. Microbiol. 36, 249–260.

9. Knutton, S., Rosenshine, I., Pallen, M. J., Nisan, I., Neves, B. C., Bain, C., Wolff, C., Dougan, G. & Frankel, G. (1998) EMBO J. 17, 2166–2176.

10. Ebel, F., Podzadel, T., Rohde, M., Kresse, A. U., Kramer, S., Deibel, C., Guzman, C. A. & Chakraborty, T. (1998) Mol. Microbiol. 30, 147–161.

11. Wilson, R. K., Shaw, R. K., Daniell, S., Knutton, S. & Frankel, G. (2001) Cell. Microbiol. 3, 753–762.

12. Sekiya, K., Ohishi, M., Ogino, T., Tamano, K., Sasakawa, C. & Abe, A. (2001) Proc. Natl. Acad. Sci. USA. 98, 11638–11643.

13. Daniell, S. J., Takahashi, N., Wilson, R., Friedberg, D., Rosenshine, I., Booy, F. P., Shaw, R. K., Knutton, S., Frankel, G. & Aizawa, S. (2001) Cell. Microbiol. 3, 865–871.

14. Ginocchio, C. C., Olmsted, S. B., Wells, C. L. & Galan, J. E. (1994) Cell 76, 717–724.

15. Reed, K. A., Clark, M. A., Booth, T. A., Hueck, C. J., Miller, S. I., Hirst, B. H. & Jepson, M. A. (1998) Infect. Immun. 66, 2007–2017.

16. Roine, E., Wei, W., Yuan, J., Nurmiaho-Lassila, E. L., Kalkkinen, N., Romantschuk, M. & He, S. Y. (1997) Proc. Natl. Acad. Sci. USA 94, 3459–3464.

17. Lucas, R. L. & Lee, C. A. (2000) Mol. Microbiol. 36, 1024–1033.

18. Lucas, R. L. & Lee, C. A. (2001) J. Bacteriol. 183, 2733–2745.

19. Darwin, K. H. & Miller, V. L. (1999) J. Bacteriol. 181, 4949–4954.

20. Eichelberg, K. & Galan J. E. (1999) Infect. Immun. 67, 4099–4105.

21. Lostroh, C. P. & Lee, C. A. (2001) J. Bacteriol. 183, 4876–4885.

22. Wei, Z., Kim, J. F. & Beer, S. V. (2000) Mol. Plant Microbe Interact. 13, 1251–1262.

23. Hendrickson, E. L., Guevera, P. & Ausubel, F. M. (2000) J. Bacteriol. 182, 3508–3516.

24. Preston, G., Deng, W. L., Huang, H. C. & Collmer, A. (1998) J. Bacteriol. 180, 4532–4537.

25. Xiao, Y. & Hutcheson, S. W. (1994) J. Bacteriol. 176, 3089–3091.

26. Xiao, Y., Heu, S., Yi, J., Lu, Y. & Hutcheson, S. W. (1994) J. Bacteriol. 176, 1025–1036.

27. Fenselau, S. & Bonas, U. (1995) Mol. Plant Microbe Interact. 8, 845–854.

28. Wengelnik, K., Van den Ackerveken, G. & Bonas, U. (1996) Mol. Plant Microbe Interact. 9, 704–712.

29. Darwin, K. H. & Miller, V. L. (2000) Mol. Microbiol. 35, 949–960.

30. Darwin, K. H. & Miller, V. L. (2001) EMBO J. 20, 1850–1862.

31. Tucker, S. C. & Galan, J. E. (2000) J. Bacteriol. 182, 2262–2268.

32. Anderson, D. M. & Schneewind, O. (1999) Mol. Microbiol. 31, 1139–1148.

33. Lloyd, S. A., Norman, M., Rosqvist, R. & Wolf-Watz, H. (2001) Mol. Microbiol. 39, 520–532.

34. Francis, M. S., Lloyd, S. A. & Wolf-Watz, H. (2001) Mol. Microbiol. 42, 1075–1093.

35. Anderson, D. M., Ramamurthi, K. S., Tam, C. & Schneewind, O. (2002) J. Bacteriol. 184, 1287–1295.

36. Evdokimov, A. G., Tropea, J. E., Routzahn, K. M., Copeland, T. D. & Waugh, D. S. (2001) Acta Crystallogr. D 57, 793–799.

37. Smith, C. L., Khandelwal, P., Keliikuli, K., Zuiderweg, E. R. & Saper, M. A. (2001) Mol. Microbiol. 42, 967–979.

38. Khandelwal P., Keliikuli K., Smith C. L., Saper M. A. & Zuiderweg E. R. (2002) Biochemistry 41, 11425–11437.

39. Birtalan, S. & Ghosh, P. (2001) Nat. Struct. Biol. 11, 974–978.

40. Stebbins, C. E. & Galan, J. E. (2001) Nature 414, 77–81.

41. Birtalan, S. C., Phillips, R. M. & Ghosh, P. (2002) Mol. Cell 9, 971–980.

42. Boyer, P. D. (1998) Biochim. Biophys. Acta 1365, 3–9.

43. Boyer, P. D. (1997) Annu. Rev. Biochem. 66, 717–749.

44. Nogi, H., Amano, T., Yoshida, M. & Kinosita, K., Jr. (1997) Nature 386, 299–302.

45. Gomis-Rüth, F. X., Moncallán, G., Pérez-Luque, R., González, A., Cabezón, E., de la Cruz, F. & Coll, M. (2001) Nature 409, 637–641.

46. Blocker, A., Gounon, P., Larquet, E., Niebuhr, K., Cabiaux, V., Parsot, C. & Sansonetti, P. (1999) J. Cell Biol. 147, 683–693.

47. Ide, T., Laarmann, S., Greune, L., Schillers, H., Oberleithner, H. & Schmidt, M. A. (2001) Cell. Microbiol. 3, 669–679.

48. Boyd, A. P., Grosdent, N., Totemeyer, S., Geuijen, C., Bleves, S., Iriarte, M., Lambermont, I., Octave, J. N. & Cornelis, G. R. (2000) Eur. J. Cell. Biol. 79, 659–671.

49. Kenny, B., DeVinney, R., Stein, M., Reinscheid, D. J., Frey, E. A. & Finlay, B. B. (1997) Cell. 91, 511–520.

50. Hayward, R. D., McGhie, E. J. & Koronakis, V. (2000) Mol. Microbiol. 37, 727–739.

51. McGhie, E. J., Hume, P. J., Hayward, R. D., Torres, J. & Koronakis V. (2002) Mol. Microbiol. 44, 1309–1321.

52. Holmström, A., Petterson, J., Rosqvist, R., Håkansson, S., Tafazoli, F., Fallman, M., Magnusson, K. E., Wolf-Watz, H. & Forsberg A. (1997) Mol. Microbiol. 24, 73–91.

53. Holmström, A., Olsson, J., Cherepanov, P., Maier, E., Nordfelth, R., Pettersson, J., Benz, R., Wolf-Watz, H. & Forsberg, A. A. (2001) Mol. Microbiol. 39, 620–632.

54. Holmström, A., Rosqvist, R., Wolf-Watz, H. & Forsberg, A. (1995) Infect. Immun. 63, 2269–2276.

55. Pettersson, J., Holmström, A., Hill, J., Leary, S., Frithz-Lindsten, E., von Euler-Matell, A., Carlsson, E., Titball, R., Forsberg, A. & Wolf-Watz H. (1999) Mol. Microbiol. 32, 961–976.

56. Dacheux, D., Toussaint, B., Richard, M., Brochier, G., Croize, J. & Attree, I. (2000) Infect. Immun. 68, 2916–2924.

57. Dacheux, D., Goure, J., Chabert, J., Usson, Y. & Attree, I. (2001) Mol. Microbiol. 40, 76–85.

58. Hartland, E. L., Daniell, S. J., Delahay, R. M., Neves, B. C., Wallis, T., Shaw, R. K., Hale, C. & Frankel, G. (2000) Mol. Microbiol. 35, 1483–1492.