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Previous Article | Next Article 
Infection and Immunity, January 2003, p. 75-85, Vol. 71, No. 1
0019-9567/03/$08.00+0 DOI: 10.1128/IAI.71.1.75-85.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.
Department of Microbiology,1 Division of Clinical Immunology and Rheumatology, University of Alabama at Birmingham, Birmingham, Alabama 352942
Received 2 July 2002/ Returned for modification 8 August 2002/ Accepted 23 September 2002
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ABSTRACT |
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INTRODUCTION |
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Pneumococcal surface protein A (PspA) is a choline-binding protein tethered to the cell surface through its C-terminal choline-binding repeat region. Previous studies demonstrated that the PspA structure consists of five distinct domains. From the N terminus to the C terminus, these domains are as follows: a signal peptide, a highly charged alpha-helical domain, a proline-rich region, a choline-binding domain, and a C-terminal 17-amino-acid (17-aa) tail (33). The alpha-helical domain contains the protection-eliciting epitopes (3) and much of the variability in sequence among different PspA proteins. The approximately 100 C-terminal amino acids of the alpha-helical domain elicit most of the cross-protective immunity (20). Based on the sequence diversity of this region, PspA proteins have been classified into two major families that consist of five clades (3, 13). The choline-binding domain, which consists of 9 or 10 20-aa repeats, is required for the attachment of PspA to the pneumococcal cell surface (18, 34, 35). Deletion of the last five repeats and the 17-aa tail results in release into culture supernatant of family 1 PspA/Rx1 (34) (protein/strain name [described below]).
PspA is required for full virulence in mouse models of pneumococcal infection (21); PspA slows the clearance of pneumococci from mouse blood (4, 21). Our laboratory and others' have shown that family 1 PspA/WU2 leads to reduced complement activation in vivo and in vitro. Tu et al. demonstrated that mice infected with WU2 maintained higher levels of C3 in their blood than mice infected with an isogenic pspA-lacking strain (28). Tu et al. also showed that the pspA-lacking strain was cleared much faster from the blood of mice. The difference in virulence of the wild-type and pspA-lacking strains disappeared when C3 or factor B knockout mice were infected. Using enzyme-linked immunosorbent assay, Abeyta et al. showed that pspA-lacking strains bound more C3 than wild-type pspA-positive strains when pneumococci were incubated with mouse serum (1). Using the bystander complement fixation assay, Neeleman et al. also showed that PspA interfered with complement activation (23). All of these results indicate that PspA may interfere with complement's host defense functions. However, all the studies described above examined PspA proteins only from PspA family 1. Analysis of sequence data showed that family 1 and family 2 PspA proteins only share approximately 40% amino acid similarity over their N-terminal alpha-helical regions (13). Thus, it was important to determine whether or not a family 2 PspA could play the same role in host-pathogen interaction.
PspA also binds lactoferrin, a multifunctional glycoprotein (10, 11, 30, 31). The exact contribution of lactoferrin binding to virulence, if any, is not yet clear. Nevertheless, the fact that virtually all pneumococci bind to lactoferrin through PspA argues for the importance of lactoferrin binding in pneumococcal pathogenesis.
In this study, we have replaced the naturally occurring family 1 PspA/WU2 with family 2 PspA/TIGR4 in the WU2 genetic background. The WU2 wild-type strain and the WU2-derived strains expressing family 2 PspA were then compared in terms of virulence, complement activation, and binding of lactoferrin.
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MATERIALS AND METHODS |
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Cloning and plasmid construction. The 3' region of the TIGR4 pspA gene was amplified by PCR using primer pair SKH73 (5'GGAAACAAGAAAACGGTATGTG3') and LSM2 (5'GCGCGTCGACGGCTTAAACCCATTCACCATTGG3'). The amplified PCR product was first cloned into pTOPO (Invitrogen, Carlsbad, Calif.), and then the insert was released from pTOPO by digestion with EcoRI and subcloned into the vector pJY4164 (34) encoding the erythromycin resistance gene. This formed plasmid pREN4. Similarly, the 5' region of the TIGR4 pspA gene was amplified by PCR using primer pair LSM12 (5'CCGGATCCAGCGTCGCTATCTTAGGGGCTGGTT3') and SKH79 (5'CCTTTAGAGTTGCATAATCATAT3'). A 500-bp fragment near the start codon of pspA of TIGR4 was produced, cloned into pTOPO, and then subcloned into pJY4164. The constructed plasmid was designated pREN3. To determine the pspA sequences from constructed strains BR92.1 and BR93.1, the pspA genes were amplified with primer pair LSM12 and LSM2 and cloned into pTOPO for sequencing.
Insertion-duplication mutagenesis. The plasmids pREN4 and pREN3 were used to transform S. pneumoniae TIGR4. Erythromycin-resistant transformants were obtained after plasmid integration via homologous recombination into chromosomes at the site of the cloned fragment. Plasmid pREN4 contains the 3' end of the pspA gene and so does not interrupt PspA expression upon insertion. This was used to link the selectable marker, erythromycin resistance, to the pspA/TIGR4 gene for later transfer of the gene to WU2. To serve as a negative control for family 2 PspA in antibody binding and lactoferrin binding experiments, a TIGR4 pspA-lacking strain was also constructed using pREN3. This plasmid had an internal portion of pspA/TIGR4 allowing the pspA gene to be inactivated by the insertion-duplication process (32). The resultant strain was designated BR61.1.
Western blot analysis. To identify each PspA, S. pneumoniae cultures were grown to exponential phase and centrifuged. The bacterial cell pellets were incubated with lysis buffer (0.01% sodium dodecyl sulfate, 0.1% sodium deoxycholate, 0.15 M sodium citrate) at 37°C for 10 min. Lysates from different strains were boiled in sodium dodecyl sulfate and electrophoresed on 10% polyacrylamide gels. After their separation, the proteins were transferred to nitrocellulose membranes and probed with rabbit antisera specific to family 1 or family 2 PspA proteins (29). The secondary antibody, goat anti-rabbit IgG (heavy and light chain)-biotin, was used in conjunction with streptavidin-alkaline phosphatase (Southern Biotechnology Associates, Inc., Birmingham, Ala.).
To examine differences in association of PspA with the bacterial surface, the presence of PspA in different culture fractions was detected by Western blot analysis as described previously (34) with some modification. Frozen bacterial stock was thawed, diluted with Todd-Hewitt broth containing 0.5% yeast extract (THY), and grown to exponential phase. To minimize contamination by PspA released from dead bacteria from the frozen stock, the exponential cultures were diluted to an optical density at 600 nm (OD600) of <0.1 with THY and then regrown to an OD600 of 0.5. One milliliter of the bacterial cell culture was centrifuged, and the supernatant was removed, passed through a 0.2-µm-pore-size filter, and collected. The bacterial cell pellet was washed with phosphate-buffered saline (PBS) (pH 7.0) and lysed with lysis buffer, and water was added to adjust to the original volume. Another 1-ml culture volume was withdrawn, and the bacterial cells were washed with PBS and then washed with 1 ml of 2% choline chloride in PBS. After centrifugation, the supernatant was collected, passed through a 0.2-µm-pore-size filter, and saved. The pellet was washed, incubated with lysis buffer, and adjusted to a 1-ml total volume with water. Equivalent amounts of all four fractions, representing 40 µl of unconcentrated cultures, were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and analyzed by Western blotting, as described above.
Detection of surface PspA. Cultures were grown in THY medium to an OD600 of 0.5, and approximately 107 CFU were withdrawn. Bacterial cells were enumerated by plating on blood agar plates. After washing with PBS, pneumococci were incubated with 100 µl of rabbit anti-PspA family-specific serum (29) (1:100 dilution with PBS) for 30 min at 37°C. After washing with PBS, the bacteria were incubated with 100 µl of fluorescein isothiocyanate-conjugated goat anti-rabbit IgG (heavy and light chain) (Southern Biotechnology Associates, Inc.) for 30 min on ice. After the final wash with PBS, bacteria were suspended in 300 µl of 2% paraformaldehyde and analyzed by flow cytometer (FACScan; Becton Dickinson, Franklin Lakes, N.J.). The results are expressed as median fluorescence intensity.
Binding of lactoferrin. Purified human iron-saturated lactoferrin (Sigma, St. Louis, Mo.) was biotinylated using the biotin-labeling kit according to the manufacturer's instructions (Boehringer Mannheim GmbH, Mannheim, Germany). Cultures were grown to OD600 = 0.5, and 107 CFU bacteria were incubated with 100 µl of biotinylated lactoferrin at various concentrations (5, 10, and 50 µg/ml in PBS) for 30 min at 37°C. Bacteria were washed with PBS and incubated with 100 µl of Alexa Fluor 488-conjugated streptavidin (10 µg/ml in PBS; Molecular Probes, Eugene, Oreg.) for 30 min on ice. The binding of lactoferrin to different pneumococcal strains is expressed as median fluorescence intensity.
Measurement of C3 deposition. To quantitate the deposition of C3 on the pneumococcal surface, the same numbers of CFU of different strains (107 CFU of culture at an OD of 0.5) were washed with PBS and incubated with 100 µl of 10% normal human serum (NHS) (Quidel, San Diego, Calif.) diluted in gelatin Veronal buffer (GVB) (Sigma) for 30 min at 37°C. Goat anti-human C3 serum (Quidel) was biotinylated as described above. After washing with PBS, the bacteria were incubated with biotinylated goat anti-human C3 antibody (1:100 dilution in PBS) for 30 min at 37°C. The bacterial suspensions were then mixed with Alexa Fluor 488-conjugated streptavidin, and finally the samples were analyzed by flow cytometry. The percentage of Alexa Fluor 488-positive bacteria (fluorescence intensity greater than 10) was used as a measure of the amount of C3 deposition.
Infection of mice. Female CBA/CaHN-XID/J (CBA/N) mice, 6 to 12 weeks old, were obtained from The Jackson Laboratories (Bar Harbor, Maine). Frozen infection stocks containing a known concentration of viable bacteria were diluted in lactated Ringer's solution to achieve the desired concentrations of bacteria. The numbers of bacteria injected were confirmed by plating on blood agar plates. For the median-time-to-death studies, CBA/N mice were challenged intravenously (i.v.) with 200 CFU of pneumococci per mouse in 200 µl of Ringer's solution. To assess the pneumococcal net growth and clearance in mouse blood in the early phase of infection, mice were challenged i.v. with 2 x 105 CFU per mouse. Blood samples were collected from the retro-orbital plexus at indicated time points. Samples were serially diluted and plated on the blood agar with, or without, erythromycin to determine the number of viable S. pneumoniae cells in the blood. When mice were coinfected with WU2 and its isogenic mutant (BR93.1 or BR92.1), 105 CFU of WU2 and 105 CFU of the mutant were mixed and injected into mice i.v. in a 0.2-ml volume. In this case, half of each blood sample was plated on blood agar, and the other half was plated on blood agar containing erythromycin. The numbers of BR93.1 and BR92.1 were determined by colony counts on the erythromycin-containing plates, and the numbers of WU2 were calculated by subtracting the numbers of CFU growing on the erythromycin plates from those growing on blood agar plates.
Sequencing and DNA analysis. DNA sequencing was completed using automated DNA sequencing (model ABI 377; Applied Biosystems, Inc., Foster City, Calif.). Sequence analyses were performed using MacVector 6.5 (Oxford Molecular, San Diego, Calif.) and Sequencer 3.0 (GeneCodes, Ann Arbor, Mich.). PspA/TIGR4 sequence data (27) were obtained from The Institute for Genomic Research website at http://www.tigr.org.
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RESULTS |
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The replacement of PspA was verified by Western blot analysis using rabbit antisera specific for family 1 and 2 PspA proteins (Fig. 1). PspA proteins from WU2 and JY1123 were recognized by anti-family 1 PspA serum, but not by anti-family 2 PspA serum. The band for cell-associated PspA/JY1123 was very faint, since most of the PspA is secreted into the culture supernatant (described below). PspA proteins from TIGR4, BR81, BR84, BR93.1, and BR92.1 were recognized by anti-family 2 PspA, but not by anti-family 1 PspA serum. PspA-negative mutants JY1119 (WU2 background) and BR61.1 (TIGR4 background) were included as controls; neither of these reacted with either anti-PspA family 1 serum or anti-PspA family 2 serum (Fig. 1).
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When strain BR92.1 was examined, an even larger fraction of PspA appeared in the supernatant and choline wash than that observed for BR93.1. Earlier reports showed that truncated PspA/Rx1 containing five choline-binding repeats, but missing the 17-aa tail, was totally secreted into culture supernatant. In contrast we observed that there was a significant amount of the mutant family 2 PspA/BR92.1 associated with the cell surface, even though this PspA contained only 4 of the normal 10 choline-binding repeats.
To further verify the presence of PspA on the bacterial cell surface in the newly constructed strains, flow cytometry was performed (Fig. 4). The median fluorescence intensities of JY1119, JY1123, and BR61.1 were comparable to those of the control sample incubated with Alexa Fluor 488-conjugated streptavidin alone. This procedure verified a lack of PspA on the surface of these strains. The median intensities of BR93.1 and BR92.1 were 51.40 and 10.55, respectively. The binding intensity of BR92.1 was only 21% of that of BR93.1 but was much greater than that of the TIGR4 PspA-negative strain, BR61.1. These results support the conclusion based on the results in Fig. 3 showing that PspA/BR92.1 is bound on the bacterial surface even though its attachment is not as robust as that of full-length family 2 PspA.
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We compared the lactoferrin binding of the family 1 (PspA/WU2) and family 2 (PspA/TIRG4) PspA proteins at three different concentrations of lactoferrin (5, 10, and 50 µg/ml) (Fig. 5). PspA-negative strains JY1119 and BR61.1 did not bind lactoferrin at any of the three concentrations (data not shown). At lactoferrin concentrations of 5 and 10 µg/ml, the lactoferrin binding of WU2 was stronger than that of TIGR4. BR93.1 showed the same result as TIGR4. The median fluorescence intensity of WU2 at a concentration of 5 µg/ml was 36.9, almost nine times as much as the medians of BR93.1 and TIGR4 (4.2 and 3.8, respectively).
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We were surprised to observe that the mutant PspA/BR92.1, missing six choline-binding repeats, did not bind lactoferrin even at a concentration of 50 µg/ml. The failure of this shorter surface-anchored PspA to bind lactoferrin was not a property of the genetic background of the strain, since both BR92.1 and BR81 showed the same result. Lactoferrin's inability to bind to BR92.1 and BR81 could not be fully explained by a lack of PspA on the cell surface, since the anti-PspA antibody detected at least 20% as much of the mutant PspA on the surface compared with the full-length family 2 PspA (Fig. 4).
C3 deposition on WU2 wild-type and mutant strains. After incubation with NHS, the amount of C3 deposition was monitored by measuring the percentage of fluorescence-positive bacteria by flow cytometry (Fig. 6). As expected, the amount of C3 that could be detected on PspA-negative strains JY1119 and JY1123 was substantially greater than the amount of C3 detected on PspA-positive strain WU2. The same amount of C3 was detected on BR93.1 as on WU2, indicating that both family 1 and family 2 PspA proteins can decrease complement activation on the WU2 surface. Deposition of C3 on BR92.1 was higher than those on BR93.1 and WU2, but less than those on JY1119 and JY1123. This observation was consistent with the relative amount of surface-associated proteins as detected by anti-PspA antibodies above (Fig. 4).
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The original studies about the attachment mechanism of PspA demonstrated that the anchoring of PspA to the bacterial cell surface requires choline-mediated interactions between the membrane-associated lipoteichoic acid and the C-terminal repeat region (35). However, there is a short and weakly hydrophobic stretch containing 13 aa crossing the last half-repeat and extending into the C-terminal 17-aa tail (Fig. 2). This sequence is highly conserved (18). It appeared to be too short to function as a membrane-spanning anchor, and Yother et al. previously reported that the elimination of the last 20 aa, which included more than half of this hydrophobic region, did not result in loss of attachment of PspA in strain Rx1 (35). It was also shown that PspA lacking the last five choline-binding repeats plus the C-terminal tail was no longer bound to the cell surface of Rx1 or WU2 (1, 34), indicating that the five remaining repeats were not sufficient for surface attachment. These results had suggested that more than five choline-binding repeats, but not the 17-aa tail, were required for surface attachment of PspA.
Our results showed that PspA/BR92.1, with four repeats, was still anchored on the cell surface of WU2 or TIGR4. However, because the deletion occurred in the internal repeat region instead of at the C-terminal end, the mutant PspA/BR92.1 still contains the last repeat and the 17-aa C-terminal tail with its hydrophobic stretch intact. Therefore, our data considered in the light of the previous studies (1, 34, 35) suggest that the C-terminal 17-aa tail can function together with the repeat units to help anchor PspA to pneumococcal surface.
We also observed that family 1 and 2 PspA proteins showed different binding activities for lactoferrin. These differences were inherited in the PspA proteins themselves, since changing genetic background did not affect the lactoferrin binding activity of PspA/TIGR4. At low concentration (5 or 10 µg/ml), family 1 PspA/WU2 binds more lactoferrin than family 2 PspA/TIGR4. It is unlikely that the different binding is due to the release of the PspA from cell surface, since both PspA proteins bound similar amounts of lactoferrin at high concentration (50 µg/ml).
Using recombinant fragments of family 1 and family 2 PspA proteins, Hankansson et al. showed that lactoferrin binds to the C-terminal end of the PspA alpha-helical region (10), the same region that elicits cross-protective immune responses (19). Alignment of the whole alpha-helical regions of family 1 PspA/WU2 and family 2 PspA/TIGR4 revealed that this family 2 PspA (418 aa) is much longer than the family 1 PspA (278 aa), and they share 32% identity overall. At the C-terminal end of the alpha-helical region where lactoferrin is thought to bind, there is only 23% identity between PspA/WU2 and PspA/TIGR4 (10, 13, 27). Thus, the disparate sequences in this region probably contribute to the different binding activity to lactoferrin. We suspect that a lactoferrin concentration of 50 µg/ml may be saturating, which therefore explains our inability to detect differential binding of family 1 and 2 PspA proteins.
It was surprising to observe that no lactoferrin binding was detected on the surfaces of BR92.1 and BR81, as the mutant family 2 PspA is missing only six choline-binding repeats and the alpha-helical region maintains intact. When the whole-cell lysate of BR92.1 was analyzed by SDS-PAGE and transferred onto nitrocellulose, the immobilized mutant PspA was found to bind lactoferrin (data not shown). Thus, the lack of binding by lactoferrin is probably not because of any change in the linear sequence of lactoferrin-binding site. Our results suggest that when PspA/BR92.1 is expressed on the cell surface, the reduction in length caused by the 121-aa deletion in the repeat region may prevent the binding site from being as accessible as it is on wild-type full-length PspA.
By incubating S. pneumoniae with normal mouse serum in vitro, both Tu et al. and Abeyta et al. observed, with Western blot or enzyme-linked immunosorbent assay methodology, that PspA-negative pneumococci bind more C3 than their isogenic PspA-positive wild-type strains (1, 28; M. Abeyta, G. G. Hardy, and J. Yother, submitted). Here, we have shown that PspA has the same inhibiting effect on deposition of human complement as was previously shown for mouse complement. In our studies, complement deposition was quantified using FACS analysis and intact living pneumococci, thus further increasing our confidence in the relevance of these findings.
The surface attachment of PspA appears to be essential for PspA to interfere with complement deposition onto pneumococci. Although PspA/JY1123 expresses the entire N-terminal functional region, the PspA is secreted and does not remain on the bacterial surface. The amount of C3 fixed on JY1123 was comparable to that of PspA-negative strain, JY1119. The amount of C3 on BR92.1 fell between the amount of C3 on BR93.1 and that on JY1119, and the difference in C3 deposition between BR92.1 and BR93.1 correlated with the results of anti-PspA antibody staining on the surface of these two strains and with the differences in virulence of these two strains. The latter observation provides the best evidence so far that the ability of PspA to interfere with complement activation plays a role in virulence.
Although the high diversity of PspA sequence would be consistent with a hypothesis that different PspA families may not have the same function, our results failed to support this idea. Family 1 and family 2 PspA proteins only share approximately 40% similarity overall, and the alpha-helical region of family 2 PspA is much longer than that of family 1 PspA. In spite of these differences, both exhibit the same virulence in a mouse bacteremia model. These similarities of our in vivo virulence results were in agreement with the virtually identical levels of in vitro complement deposition on BR93.1 and WU2.
These results suggest that these two PspA proteins, representing the two major PspA families, exert the same negative effects on complement deposition. Because of the sequence variation in PspA, it is likely that the ability of PspA proteins to interfere with complement deposition may depend upon a shared area of conformations of family 1 and 2 PspA proteins rather than upon the specific conservation of a linear amino acid sequence. Our present findings are consistent with a hypothesis that the major effect of different PspA proteins on virulence is mediated by their common ability to inhibit complement deposition on pneumococci.
Lactoferrin is a multifunctional protein which plays an important role in innate immunity. It is cytotoxic for bacteria and inhibits bacterial adherence and colonization on mucosal surfaces (2, 8). Importantly, it was also reported that lactoferrin affects the activation of the classical complement pathway (15, 16, 25). In our studies, family 1 and family 2 PspA proteins did not show the same binding activity to lactoferrin at low concentrations (5 or 10 µg/ml), which are near the lactoferrin concentrations in NHS. However, they showed the same effects on complement activation. Moreover, the truncated PspA/BR92.1 did not bind to lactoferrin but still maintained the anticomplementary function in human serum and was still partially virulent in mice. These results may indicate that the function of inhibiting human complement deposition probably does not depend on the PspA binding with serum lactoferrin. The results suggest, but do not prove, that lactoferrin may not play an important role in the pneumococcal bacteremia or sepsis model. It must be remembered that our lactoferrin binding studies were done with human lactoferrin sharing only 69% sequence identity with mouse lactoferrin. Thus, based on these data, we cannot completely exclude an important role of the PspA-human lactoferrin binding in bacteremia or sepsis in humans. The PspA-lactoferrin interaction may play an important role in nasopharyngeal colonization, which is important for both pneumococcal acquisition and spread to other humans.
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ACKNOWLEDGMENTS |
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This work was supported by NIH grant AI42183 (A.J.S.) and by NIH grant AI21548 and the Carsten Cole Buckley Pediatric Meningitis Research Fund (D.E.B.).
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FOOTNOTES |
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