Vaccine Development Laboratory, National Public Health Institute, Mannerheimintie 166, FIN-00300, Helsinki, Finland
Correspondence
Vesa P. Kontinen
vesa.kontinen{at}ktl.fi
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ABSTRACT |
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Present address: Institute of Biotechnology, Biocentre 1, PO Box 56, FIN-00014 University of Helsinki, Helsinki, Finland.
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INTRODUCTION |
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In the Gram-positive Bacillus subtilis, the cell membrane is surrounded by a matrix of cell wall polymers, peptidoglycan and anionic teichoic (or teichuronic) acids (reviewed by Archibald et al., 1993). The matrix polymers form a porous structure that allows macromolecules up to a molecular mass of about 50 kDa (Demchick & Koch, 1996
) to pass through the wall. Teichoic acids, which are polymers of glycerol (or ribitol) phosphate units, are attached either to the peptidoglycan (wall teichoic acids) or to the cell membrane (lipoteichoic acids). These anionic polymers are abundant constituents of the wall and provide a high density of negative charge on the cell envelope and a high capacity to bind divalent metal ions and other cationic molecules (Beveridge & Murray, 1980
; Peschel et al., 1999
; Petit-Glatron et al., 1993
). The negative charge as well as the wall's ability to bind cations is reduced by D-alanine ester substitution of teichoic acids (Hyyryläinen et al., 2000
; Perego et al., 1995
). Since the B. subtilis cell does not have a membrane-enclosed periplasm, secretory proteins move directly through the cell wall into the external medium. However, the pore size and negative charge of the wall matrix can be expected to limit the protein traffic (Demchick & Koch, 1996
; Merchante et al., 1995
). Some proteins also remain in the matrix of the cell wall. Among them are WprA protease, wall-associated protein WapA and autolysins (Margot & Karamata, 1996
; Smith et al., 2000
; Yoshida et al., 1995
).
Secreted proteins emerge from the translocase to the compartment between the cell wall and the cytoplasmic membrane. This is a demanding environment for protein folding owing to the high density of negative charge, high concentration of cations and low pH immediately outside the membrane. These factors most likely pose stringent requirements for the folding kinetics of secreted proteins. Native proteins compatible with the conditions at the membranewall interface fold with fast kinetics into their normal conformation. In contrast, heterologous proteins, produced in B. subtilis for biotechnical applications, may have slow folding kinetics. Furthermore, they are usually more susceptible to proteolytic degradation than native proteins (Stephenson et al., 1998). The incompatibility associated with heterologous proteins may also result in misfolding and aggregation (Bolhuis et al., 1999
; Meens et al., 1993
). There are regulatory mechanisms (CssRS two-component system) to sense the accumulation of misfolded proteins at the membranewall interface and to activate the synthesis of HtrA-type cleaning proteases' (Hyyryläinen et al., 2001
). This is thought to be one of the major reasons for the low production levels of heterologous proteins in industrial applications.
The major extracytoplasmic folding factor in B. subtilis is the PrsA protein, which belongs to the parvulin family of PPIases (Rahfeld et al., 1994). PrsA is a typical bacterial lipoprotein anchored to the cytoplasmic membrane by an N-terminal diacylglyceryl moiety (Kontinen & Sarvas, 1993
; Kontinen et al., 1991
; Leskelä et al., 1999b
). The hydrophilic, positively charged protein domain is located on the outer surface of the membrane, as indicated by its accessibility to external trypsin in protoplasts (Leskelä et al., 1999b
). Our data showed a linear relationship between the amount of cellular PrsA and the secretion of overproduced AmyQ
-amylase into the culture medium (Vitikainen et al., 2001
). The PrsA protein does not influence either the expression or the translocation of secretory proteins, but it is required for their folding and stability in the post-translocational phase of secretion at the membranecell wall interface (Hyyryläinen et al., 2000
, 2001
; Jacobs et al., 1993
; Leskelä et al., 1999b
; Vitikainen et al., 2001
). PrsA is an essential cell component, suggesting that it also affects the folding and stability of some essential proteins involved in the synthesis of the cell wall or the function of the cell membrane (Vitikainen et al., 2001
).
This study was designed to elucidate the role of the cell wall for the function of the PrsA folding factor. To accomplish this, we developed a method to pulsechase label protoplasts. We compared the secretion kinetics and stability of a model protein, AmyQ -amylase, in protoplasts and rods. The results indicated that unlike in rods, PrsA is not needed for the folding, stability or secretion of AmyQ in protoplasts. Furthermore, in the absence of the wall, a substantial fraction of AmyQ remains cell-associated.
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METHODS |
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Preparation of protoplasts.
Cells overproducing B. amyloliquefaciens -amylase (AmyQ) were grown in SMSa minimal medium up to the late exponential growth phase. In this growth phase, the culture was typically at a cell density of 50100 Klett units (OD600 of about 0·51·0), and the cells efficiently secreted AmyQ. In a typical experiment, cells from 20 ml culture were harvested and resuspended in 2 ml SMSb [SMSa with 20 % (w/v) sucrose and 10 mM Mg2+ for the stability of protoplast membrane] containing lysozyme (1 mg ml-1) and mutanolysin (100 U ml-1) to remove the cell wall. The lysozyme and mutanolysin were purchased from Sigma. The cell suspension was slowly shaken at 37 °C and the conversion to protoplasts was monitored by phase-contrast microscopy. After incubation for 30 min, all the cells had turned into protoplasts, as revealed by phase-contrast microscopy. Electron microscopy showed that the total surface of the cytoplasmic membrane was exposed, with no visible structures of the cell wall remaining (Fig. 1
A). Protoplasts were harvested by centrifugation at 9000 g for 4 min and resuspended in 3 ml SMSb. Electron microscopy was performed as described previously (Lounatmaa, 1985
).
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Other methods.
The trypsin sensitivity of AmyQ secreted from protoplasts was determined as follows. A series of protoplast preparations in 3 ml SMSb each were prepared as described above, followed by incubation at 37 °C for 10 min with shaking (100 r.p.m.) and then continued for 1 h in the presence of different concentrations (0·2, 1, 5, 25 and 100 µg ml-1) of trypsin. After the incubation, samples (0·4 ml) were filtered through Millex-LCR13 filter units (Millipore), trypsin inhibitor was added at a concentration exceeding that of trypsin by at least twofold and the -amylase activities of the samples were determined. The trypsin sensitivity of PrsA and PrsA3 in protoplasts was determined with 50 µg ml-1 trypsin in a similar way to that of AmyQ, except that the PrsA content of the protoplast samples was analysed by immunoblotting.
Extracellular -amylase activity was determined using Phadebas Amylase Test Tablets (Pharmacia) as described previously (Leskelä et al., 1999a
).
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RESULTS |
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We next examined the incorporation of metabolic label into the protoplasts. These were labelled with 50 µCi [35S]methionine for 45 s, then chased with non-radioactive methionine for 10 min, followed by separation of the protoplast and supernatant fractions by centrifugation. About 35 % of the added label was incorporated into the protoplasts, as determined by counting radioactivity of TCA precipitates immediately after labelling. This efficiency of label incorporation was only slightly less than that observed with rods (45 %) labelled under identical conditions. SDS-PAGE and fluorography revealed that the protoplasts were metabolically active: the labelling of proteins was effective and comparable to that in the rods (Fig. 1B). To determine the stability of the protoplasts during the procedure of pulsechase, we scanned densitometrically one strongly labelled putative intracellular protein (indicated with an arrow in Fig. 1B
). In the non-chased sample, about 7 % of this protein resided in the supernatant fraction after pelleting of protoplasts most likely due to breakage of some of the protoplasts during the separation of the protoplast pellet and supernatant fractions subsequent to the pulsechase labelling (Fig. 1B
). This amount did not increase during the chase of up to 40 min (Fig. 1B
and data not shown), indicating that the stability of the protoplasts did not deteriorate during the pulsechase labelling. The pattern and intensity of other protein bands also remained unchanged during the chase. No cellular proteins were found in the rod supernatant.
Cell wall matrix influences the kinetics of the signal peptide processing
The AmyQ proteins radiolabelled during a pulse were immunoprecipitated by specific antibodies. The immunoprecipitated preAmyQ and AmyQ polypeptides were separated in SDS-PAGE and visualized by fluorography. In a parallel experiment, rods of the same strain were pulsechase labelled and analysed similarly. Similar amounts of labelled and immunoprecipitated AmyQ proteins were found in the prechase preparations of rods and protoplasts (Fig. 1C). In rods, the cleavage of the signal peptide was completed within 2 min of the chase. Mature
-amylase was chased from cells and accumulated in the medium (Fig. 1C
). In protoplasts, the signal peptide processing was clearly slower. After 2 min of chasing, less than 50 % of the precursor synthesized during the pulse was chased into the mature form and even after 10 min of the chase, some preAmyQ was still detected (Fig. 1C
). Mature AmyQ, but not the precursor, was released into the medium, although the amount was only about 15 % of that released from the rods. The proportions of the intensities of the AmyQ bands varied to some extent from one experiment to another (see Fig. 2
B). The processing and export of AmyQ were similarly inhibited in both the rods and protoplasts by sodium azide, an inhibitor of the SecA ATPase of the protein translocator (Fig. 1C
). Thus, not only are the protoplasts metabolically active, but also they secrete proteins into the medium through the Sec-dependent pathway.
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DISCUSSION |
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In rods, nascent AmyQ most likely interacts with the negatively charged polymers in the cell wall and on the cytoplasmic membrane, consequently resulting in AmyQ misfolding that renders it susceptible to the extracytoplasmic quality-control proteases activated by the ensuing folding stress (Hyyryläinen et al., 2001). A likely role of PrsA is to prevent such interactions and thus facilitate the folding. This function may be a chaperone-like function rather than being due to PPIase activity. There is no clear saturation of the PrsA-enhanced AmyQ secretion, when PrsA is produced in high excess as compared with AmyQ (Vitikainen et al., 2001
), suggesting that the crucial mechanism of PrsA in AmyQ secretion is a chaperone-like function. Furthermore, the B. licheniformis
-amylase AmyL, which is like AmyQ secreted in a PrsA-dependent manner (Kontinen & Sarvas, 1993
and unpublished results) does not contain any prolines in the cis conformation (PDB code: 1BLI), again pointing to a non-PPIase mechanism in the PrsA-catalysed
-amylase secretion.
Many PPIases have, in addition to their prolyl isomerase activity, a chaperone-like activity. This has been shown with all three classes of PPIases, cyclophilins, FK506-binding proteins and parvulins (Arié et al., 2001; Behrens et al., 2001
; Bose et al., 1996
; Freeman et al., 1996
; Freskgard et al., 1992
; Ramm & Plückthun, 2000
; Scholz et al., 1997
). The PPIase activity-independent chaperone activity is located either in the PPIase domain, as in the E. coli FkpA protein (Ramm & Plückthun, 2001
), or outside it, as in the E. coli SurA protein (Behrens et al., 2001
).
A major effect of the removal of the cell wall was to impair drastically the overall secretion of AmyQ. Significantly, the pulsechase labelling revealed that a substantial fraction of mature AmyQ (50 %) remained firmly associated with the protoplast membrane, whereas in rods, AmyQ was completely released into the external medium. We can envisage several possible mechanisms for the defective release of AmyQ from the protoplast membrane. There may be in the cell wall a releasing factor that facilitates AmyQ release from the membrane. Its absence in protoplasts could be responsible for the observed retention of AmyQ in the membrane. This operational factor might be a cell-wall-associated protein(s), wall-bound divalent metal ion(s) or the cell wall matrix itself. Retardation in protoplasts of another secreted protein, AmyE of B. subtilis, has been described previously, and a mechanism coupled with folding specifically dependent on calcium ions was suggested (Haddaoui et al., 1997
; Leloup et al., 1999
). However, the calcium-enhanced release is most likely not a general release mechanism in the secretion of proteins since, in our study, the secretion of AmyQ from protoplasts was found to be independent of calcium. Consistent with this conclusion, it has been shown previously that divalent cations facilitate the production of enzymically active protease by protoplasts of B. amyloliquefaciens, whereas they have no effect on the production of
-amylase (Sanders & May, 1975
). We could not determine whether Mg2+ could replace Ca2+, because of the fragility of the protoplasts in the absence of Mg2+.
As a negatively charged matrix, the cell wall may facilitate protein release from the membrane by electronegative interactions. Alternatively, removal of the cell wall may perturb functions of the cell membrane such as proton motive force, protein translocation, signal peptide cleavage or expression of proteases associated with the wall and wallmembrane interface and thereby indirectly affect the AmyQ release from the membrane. We observed a moderate defect of the signal peptide processing, but this is hardly the reason for the AmyQ retention, since mutants that are defective in the signal peptide processing such as ecs (Pummi et al., 2002) do not cause retention of mature AmyQ in the cell membrane. The high metabolic activity of the protoplasts in terms of protein synthesis excludes major dysfunctions of the membrane. Our results also indicated that PrsA is not involved in the release mechanism.
The pulsechase-labelling method of protoplasts described above can be used to study late stages of protein secretion in B. subtilis by making the components involved accessible to external manipulation and addressing the role of the cell wall matrix. The main problem of the method is the considerable lysis of protoplasts during their preparation and handling. This is, however, mitigated by their stability during pulsechase labelling. Another limitation is the inability of lysozyme to remove all components of the cell wall; the protoplasts most likely still contain membrane-bound lipoteichoic acids and precursors of the peptidoglycan, which, in the absence of the rest of the wall matrix, may present non-physiological functions. Lysozyme may partially neutralize the negative charge on the protoplast surface and thereby influence the properties of the protoplasts.
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ACKNOWLEDGEMENTS |
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Received 7 February 2002;
revised 27 September 2002;
accepted 18 November 2002.