1 Clinical Dental Science, University of Liverpool, Liverpool L69 3GN
2 Department of Medical Microbiology and Genito-Urinary Medicine, University of Liverpool, Liverpool L69 3GN
3 School of Chemical and Life Science, The University of Greenwich, Chatham Maritime Campus, Pembroke, Chatham ME4 4TB
Correspondence
John W. Smalley
josmall{at}liv.ac.uk
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ABSTRACT |
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INTRODUCTION |
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METHODS |
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Preparation of iron(III) protoporphyrin IX solutions.
Iron(III) protoporphyrin IX was prepared as either the monomer, Fe(III)PPIX.OH (haematin), or the µ-oxo oligomer, [Fe(III)PPIX]2O, as follows. Bovine haemin [Fe(III)PPIX.Cl] was dissolved in 0·14 M NaCl in 0·1 M Tris, at pH 9·8, to give a 1 mM stock solution. The pH was then adjusted to pH 7·5 by slow drop-wise addition of dilute HCl. This was further diluted with 0·14 M NaCl, 0·1 M Tris/HCl, at pH 7·0, to give a 100 µM solution containing a mixture of both the µ-oxo oligomer, or in NaCl/phosphate at pH 6·5 to yield a solution comprising predominantly the monomeric iron(III) protoporphyrin IX species (Silver & Lukas, 1983; Miller et al., 1987
). The UV-visible spectra of these solutions were then recorded to confirm the presence of the monomeric and µ-oxo oligomeric forms of iron(III) protoporphyrin IX, which can be seen by the presence of prominent Soret bands at 365 and 385 nm, respectively (Silver & Lukas, 1983
). The solutions were used immediately and no longer than 1 h after preparation.
Reaction of whole cells with iron(III) protoporphyrin IX species
(i) Spectrophotometic titrations of cells with iron(III) protoporphyrin IX.
Suspensions of cells (1 ml) standardized to an OD400 of 0·5 in NaCl/Tris, pH 7·0, or in NaCl/phosphate, pH 6·5, were used. For each haemcell titration four 1 cm path length semi-micro optical cuvettes (AD) were set up as follows: A and B contained 1 ml of the above cell suspension, and C and D contained 1 ml of appropriate buffer. To cuvettes A and C, 10 µl iron(III) protoporphyrin IX solutions (100 µM on a haem monomer basis) at pH 6·5 or 7·0, were added. These titrations were carried out at pH 7·0 to examine the interactions of cells with a mixture of both Fe(III)PPIX.OH monomers and µ-oxo oligomers, and at pH 6·5 to examine the interaction with Fe(III)PPIX.OH monomers in the absence of µ-oxo oligomers (Silver & Lukas, 1983; Miller et al., 1987
). Cuvettes B and D received 10 µl aliquots each of the appropriate buffer. The contents of the cuvettes were mixed and the UV-visible spectra recorded immediately. Further sequential 10 µl additions of either iron porphyrin or buffer were made and spectra were recorded immediately after mixing. The spectra for cuvettes A and C were corrected by subtraction of spectra of the cells alone (cuvette B) or buffer (cuvette D). Difference spectra were obtained by subtracting the corrected spectrum obtained for the haemcell interaction from that of the corrected haem spectrum i.e. (A-B)-(C-D). Where appropriate, the overall absorbance differences (
A) in each spectrum between the minimum and maximum points were plotted versus the concentration of iron porphyrin added to the cell suspension.
(ii) Time-course of interaction of cells with iron(III) protoporphyrin IX solutions.
Cell suspensions standardized as above were mixed with 25 nmol Fe(III)PPIX.OH in a total volume of 1 ml NaCl/phosphate, pH 6·5, at 20 °C and the spectrum was recorded immediately, and then at 10 min intervals. These spectra were corrected for the background absorbance due to the presence of cells in suspension.
(iii) Exposure of whole cells to iron(III) protoporphyrin IX and identification of haem-binding components.
For these experiments cells were firstly grown on M9 Minimal Salts Medium agar and suspensions of these (0·5x109 in 0·5 ml) were incubated at 37 °C for 30 min with an equal volume of concentrations of either Fe(III)PPIX.OH or [Fe(III)PPIX]2O (0160 µM on a haem monomer basis) in 0·14 M NaCl buffered either at pH 6·5 or at pH 7·5, respectively. The incubation mixtures were centrifuged at 13 000 g for 5 min, and the pelleted cells were washed three times in the original assay buffer to remove any residual unbound iron porphyrin. The cell pellets were resuspended in 1 ml of the above buffers and solubilized at 37 °C in non-reducing application buffer for subsequent SDS-PAGE and staining with tetramethylbenzidine/H2O2 for detection of haem protein-associated peroxidase activity as described previously (Smalley et al., 2001). The equivalent of 8x107 solubilized cells were loaded per track. Following SDS-PAGE and tetramethylbenzidine/H2O2 staining, the gels were counter-stained for protein with 0·1 % (w/v) Coomassie blue in 50 % (v/v) methanol, 7 % (v/v) acetic acid and 43 % H2O, and diffusion destained in the above solvent mixture, to allow a precise identification of the tetramethylbenzidine/H2O2-positive polypeptides.
Iron(III) protoporphyrin binding assays.
Iron protoporphyrin IX binding to whole cells was performed at pH 7·5 to examine the binding of the µ-oxo oligomer which exists as the predominant ferrihaem species at this pH (Silver & Lukas, 1983; Miller et al., 1987
). For these experiments, as with the cellhaem titrations, cells grown on M9 Minimal Salts Medium agar were used since this medium contains no endogenous haem which could have interfered with the iron(III) protoporphyrincell binding interactions. Cells were suspended in NaCl/Tris, pH 7·5, standardized to 109 ml-1 and aliquots (0·5 ml) were mixed with 0·5 ml NaCl/Tris buffer (as above) containing iron(III) protoporphyrin IX to give a range of concentrations between 0 and 240 nmol ml-1 (on a haem monomer basis). These were incubated at 37 °C for 30 min by end-over-end mixing. The incubation mixtures were centrifuged at 11 000 g for 10 min at 20 °C and the pelleted cells were washed and recentrifuged three times in the same buffer to remove any unbound residual iron porphyrin. The cell-bound iron(III) protoporphyrin IX was converted to the iron(II) form by the addition of freshly prepared Na2S2O4 (10 mM final molarity) and assayed as the pyridine-haemochrome as described previously (Smalley et al., 2001
). The amounts bound to the cells were expressed on a haem monomer basis.
Outer membrane extraction.
The outer membrane fraction was isolated from cells grown on M9 Minimal Salts Medium agar using the EDTA-shearing method as described previously (Smalley et al., 2001).
Catalase assays.
Catalase activity was determined using the UV absorbance method of Beers & Sizer (1952) as described previously (Smalley et al., 2000
). Cell suspensions (0·5x109 in 1 ml) were exposed to iron(III) protoporphyrin IX monomers or µ-oxo oligomers (60 nmol ml-1, on a monomer basis) at either pH 6·5 or 7·5, respectively, for 30 min at 37 °C as above, and washed repeatedly to remove any unbound iron porphyrin. Suspensions of these cells (0·5x108 ml-1) carrying bound iron(III) protoporphyrin IX were mixed with H2O2 (10 mM final molarity) in the appropriate buffer and the UV absorbance at 240 nm monitored over the first 60 s of the reaction at 20 °C. The activities of the cells with bound iron porphyrin were corrected for endogenous background catalase activity. The rates of H2O2 destruction were calculated by regression analysis from the initial linear decrease in A240 using GraphPad Prism.
Spectrophotometry.
UV-visible spectra were recorded in an Ultrospec 2000 scanning spectrophotometer (Pharmacia Biotech) in quartz or plastic 1 ml semi-micro cuvettes (Elkay UltraVu) with a 1 cm path length.
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RESULTS |
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DISCUSSION |
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To understand the behaviour of iron(III) protoporphyrin IX it is necessary to appreciate that these molecules exist in both monomeric and µ-oxo oligomeric forms. In solution, the formation of the [Fe(III)PPIX]2O complex from Fe(III)PPIX.OH monomers is promoted in the presence of base, and this complex can dissociate in the presence of protons according to the equation:
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The pH of the surface liquid layer in the normal lung is reported to be around 6·9 (Jayaraman et al., 2001). For this reason, the interaction of bacterial cells with iron(III) protoporphyrin IX was initially studied at pH 7·0. At neutral pH, free ferrihaems will be present as a mixture of both the monomeric and µ-oxo oligomeric forms as a result of the pH-dependent equilibrium. Under these conditions B. cepacia genomovar IIIa strains mediated the depletion of the monomer in solution and the formation of [Fe(III)PPIX]2O. This effect was not observed for the genomovar I strains. To our knowledge the ability to mediate this transformation is a novel phenomenon and has not been described in any other biological systems, although the [Fe(III)PPIX]2O complex, which is the major haem species in the green-black pigment of P. gingivalis (Smalley et al., 1998
), is generated as a result of degradation of both oxy- and deoxyhaemoglobin (Smalley et al., 2002
). Thus, the pH of the liquid surface layer of the lung in health and disease will have a major influence on the behaviour of any free ferrihaems. It is noteworthy that endobronchial pHs of between 6·58 and 6·62 have been recorded in individuals with chronic lung disease and Gram-negative pneumonia (Bodem et al., 1983
). In view of this, experiments were also conducted at pH 6·5 to permit the interaction of cells with Fe(III)PPIX.OH monomers in the absence of the µ-oxo oligomeric form. Under these conditions the genomovar IIIa strains were also able to achieve the transformation of Fe(III)PPIX.OH into [Fe(III)PPIX]2O. These findings are important since they show that genomovar IIIa strains would be able to generate the µ-oxo oligomer from any free iron(III) protoporphyrin IX monomers both in the healthy lung (at neutral pH) and under the slightly lower pH conditions which may prevail during chronic lung inflammation. These observations may explain why epidemic genomovar IIIa strains are frequently found able to superinfect lungs already colonized by other species, including Pseudomonas aeruginosa and other B. cepacia genomovars (Hart & Winstanley, 2002
; Mahenthiralingam et al., 2001
) where, in the presence of chronic inflammation, the pH is likely to be lower than neutral (Bodem et al., 1983
).
Difference spectra obtained from titrations of iron(III) protoporphyrin IX with cells of the genomovar IIIa strains at pH 7·0 revealed that there was a continuous formation of [Fe(III)PPIX]2O from Fe(III)PPIX.OH monomers. Moreover, the alkaline growth end point generated by all the strains in this study (as demonstrated by the production of pink colonies on Burkholderia cepacia Medium) is significant since this would favour µ-oxo oligomer formation from any free Fe(III)PPIX.OH (Silver & Lukas, 1983) and encourage haem aggregation on the cell surface. For these reasons it was felt pertinent to assess the cellular binding capacity for iron(III) protoporphyrin IX in the µ-oxo oligomeric form. The genomovar IIIa strains BC7 and C5424 displayed greater binding capacities for [Fe(III)PPIX]2O than the genomovar I isolates (LMG 17997 and ATCC 25416), and Scatchard plots indicated monophasic binding. The apparent affinity constants were of the order 104 M-1. Since we were unable to demonstrate a bathochromic shift of the Soret band during the titrations of cells at the lowest concentrations of added iron(III) protoporphyrin IX at pH 7·0 (other than an increase in A385 indicative of [Fe(III)PPIX]2O formation) we suggest that these curves represent binding to the cell surface via aggregation, rather than to some specific receptor for the µ-oxo oligomer. This is supported by our observation that the Soret bands resulting from incubation of cells of genomovar IIIa strains BC7 and C5424 with a fixed concentration of iron(III) protoporphyrin IX were both decreased in intensity and broadened, features indicative of aggregation. Aggregation of newly generated [Fe(III)PPIX]2O and its subsequent removal from solution and deposition on the cell surface would result in a further shift in the equilibrium towards production of the µ-oxo oligomeric species from any monomers remaining in solution and encourage further deposition of µ-oxo oligomer molecules on the cell surface. It is thus noteworthy that genomovar IIIa strains BC7 and C5424 yield green-brown colonies when grown on blood agar and that incubation of oxyhaemoglobin with these strains results in the generation of a Soret band absorbing component with features of the [Fe(III)PPIX]2O complex (Smalley et al., 2002
; unpublished data). Although we cannot rule out the possibility that these isolates produce pyoverdin-like molecules, our observations support the contention that B. cepacia can generate and bind [Fe(III)PPIX]2O during growth on blood-containing media. However, it is not yet clear as to whether other cell-surface components bind [Fe(III)PPIX]2O. Thus, the ability to mediate the transformation of the Fe(III)PPIX.OH monomers into [Fe(III)PPIX]2O is a novel phenomenon associated with genomovar IIIa isolates and constitutes a potential virulence determinant in this group of organisms.
Tetramethylbenzidine/H2O2 staining of cellular components on SDS-polyacrylamide gels after binding of iron(III) protoporphyrin IX has been used to identify cell-surface HBPs (Lee, 1992; Mazoy & Lemos, 1996
; Smalley et al., 1993
; Stugard et al., 1989
). When genomovar IIIa strains were exposed to increasing concentrations of iron(III) protoporphyrin IX we observed the dose-dependent binding of iron(III) protoporphyrin IX monomers to two outer-membrane proteins of 77 and 149 kDa, which were not found in the genomovar I strains even after exposure to the highest concentrations of either Fe(III)PPIX.OH or [Fe(III)PPIX]2O. In contrast to the 97 kDa putative HBP, the 77 and 149 kDa proteins were only stained after prior exposure to iron(III) protoporphyrin IX. We attribute the low degree of tetramethylbenzidine/H2O2 staining at pH 7·5 to the binding of some Fe(III)PPIX.OH monomers which would be present in the µ-oxo oligomer haem solution as a result of the equilibrium between the two forms. Thus, the fact that 77 and 149 kDa proteins were stained at 6·5 and not at 7·5 indicated that these components preferentially pick up monomers rather than µ-oxo oligomers. Since Fe(III)PPIX.OH monomers are more catalytic than the µ-oxo oligomers in their ability to destroy hydrogen peroxide (Brown et al., 1970
; Jones et al., 1973
), it is likely that Fe(III)PPIX.OH molecules bound to the cell surface at pH 6·5 would more efficiently protect against H2O2 than µ-oxo oligomers. However, it should be noted that the µ-oxo oligomer (Brown et al., 1970
) may be preferentially oxidized and partially decomposed compared to the monomer during exposure to H2O2 (Grinberg et al., 1999
) and thus constitute a cell-surface layer which can act as a sacrificial barrier against this oxidant as reported for P. gingivalis (Smalley et al., 2000
).
We have previously shown that the 97 kDa putative HBP appears to pick up both Fe(III)PPIX.OH monomers and µ-oxo oligomers (Smalley et al., 2001). However, importantly, we found in this study that, compared to the 149 and 77 kDa proteins, the 97 kDa putative HBP was stained even in the absence of any exogenously added haem and was not stained using the peroxidase substrate tetramethylbenzidine in a dose-dependent fashion after exposure of the cells to iron protoporphyrin IX. We conclude from this that this protein may contain an endogenously synthesized haem-like prosthetic group and that it may not have a true haem-binding function.
Although Fe(III)PPIX.OH monomers react spontaneously in solution to give the µ-oxo oligomeric species (Silver & Lukas, 1983; Miller et al., 1987
) (through the reverse of reaction 1), we propose that the 149 and 77 kDa proteins act together or separately as templates to facilitate the conversion of the Fe(III)PPIX.OH into [Fe(III)PPIX]2O. The continuous production would only occur if the 149 and 77 kDa protein receptor sites for the monomer were not saturated with the newly formed [Fe(III)PPIX]2O. It is not clear what bonding interactions take place between the ferrihaem monomers and these outer-membrane proteins, but we propose the following mechanism to account for the formation of [Fe(III)PPIX]2O from Fe(III)PPIX.OH as a continuous process. This is based upon the known behaviour of iron porphyrins which form µ-oxo oligomers (Silver & Lukas, 1983
; Miller et al., 1987
). First, monomers may initially become bonded to the NH2 or COOH groups of these proteins either via the haem iron (by replacement of the weakly axially bonded H2O in the sixth co-ordinate position), or via a peripheral substituent on the iron porphyrin (e.g. the methyl, vinyl or carboxylate groups). Formation of the Fe-O-Fe bond of the [Fe(III)PPIX]2O complex could then occur through the reaction of the bound monomer with another Fe(III)PPIX.OH molecule (via the hydroxyl groups; Silver & Lukas, 1983
), itself also bonded to the protein, or free in solution. Bonding of the iron protoporphyrin IX monomers to the protein(s) would facilitate this process since it would encourage electron withdrawal at the periphery of the porphyrin ring, a phenomenon which is known to encourage formation of the µ-oxo-bridged structure (Miller et al., 1987
). This would weaken any peripheral or axial bonds and force the release of the µ-oxo oligomer from the protein(s). This reaction would be encouraged at a slightly alkaline pH and aggregation of the newly formed and released [Fe(III)PPIX]2O molecules would then be possible.
The inability of the 77 and 149 kDa proteins to bind [Fe(III)PPIX]2O molecules (as evidenced by the lack of TMB/H2O2 staining at pH 7·5) indicates that the µ-oxo oligomeric molecules may not be retained on the protein(s) once they have been formed. Indeed, this is supported by the spectroscopic data which show that [Fe(III)PPIX]2O molecules, once generated, remain (initially at least) in solution. The above data also suggest that the 77 and 149 kDa proteins may not just represent a surface seed for the deposition of the µ-oxo oligomers on the cell surface, although we cannot rule out the possibility that some other component may also be responsible for mediating [Fe(III)PPIX]2O formation and cell-surface binding. Importantly, however, aggregation of the µ-oxo oligomers would facilitate the shift in the equilibrium towards the production of more of the µ-oxo oligomeric species from the monomer due to its removal from solution.
When cell-ferrihaem titrations were conducted with Fe(III)PPIX monomers in the absence of [Fe(III)PPIX]2O, genomovar IIIa strain BC7 mediated a shift in the equilibrium towards the production of [Fe(III)PPIX]2O, although this effect was not as pronounced as at the higher pH. Moreover, at pH 6·5 there was evidence from the difference spectra that the cells became saturated over the same concentration range studied. At this pH the equilibrium would favour the existence of the monomeric form and it is likely that under these conditions the 149 and 77 kDa proteins would remain occupied by Fe(III)PPIX.OH molecules. As the µ-oxo oligomer has a greater propensity to aggregate than the monomeric form, binding and conversion of Fe(III)PPIX.OH into [Fe(III)PPIX]2O would provide an advantage by forming a protective barrier against ingress of reactive oxidant species to the cell surface. In addition, bound µ-oxo oligomers would be able to serve defensively by destroying peroxide through their inherent catalase activity. It has been reported previously that strains of B. cepacia display catalase activity (Gessner & Mortensen, 1990; Chester, 1979
; Lefebre & Valvano, 2001
). However, it was found that the catalase activities of the two genomovar IIIa isolates were between 5- and 10-fold greater than those of the two genomovar I strains after exposure and binding of the monomer and µ-oxo oligomer (after correction for intrinsic cellular catalase activity). In addition, the catalase activities of cells of both genomovar IIIa and I strains after exposure to iron(III) protoporphyrin IX monomers were double those of cells carrying bound µ-oxo oligomers. This finding is predictable on the basis that the catalase activity of the monomer is higher than the µ-oxo oligomer (Jones et al., 1973
; Brown et al., 1970
). The increased catalase activity displayed by the genomovar IIIa isolates as a result of binding of ferrihaems would provide an advantage over other genomovars to endure fluxes of H2O2 released from neutrophils. Such iron porphyrin binding behaviour may aid colonization and establishment of infections of the CF lung caused by this important group of respiratory pathogens. Examination of the SDS-PAGE profiles of the strains described in our previous paper (Smalley et al., 2001
) revealed that proteins with molecular masses of 77 and 149 kDa were only present in genomovar IIIa strains. Given the role of these cell-surface proteins in picking up haems, it is possible that they may be targeted specifically to inhibit cellular haem-binding activity. Alternatively, as they appear to be prominent components on the bacterial surface, they might be used as suitable candidates for vaccine development.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 29 November 2002;
revised 17 January 2003;
accepted 17 January 2003.
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