School of Biosciences, The University of Birmingham, Edgbaston, Birmingham B15 2TT, UK1
Research and Technology, BNFL, Springfields Works, Preston PR4 OXJ, UK2
Author for correspondence: Lynne E. Macaskie. Tel: +44 121 414 5889. Fax: +44 121 414 6557. e-mail: l.e.macaskie{at}bham.ac.uk
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ABSTRACT |
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Keywords: Citrobacter, lipopolysaccharide, phosphatase, metal biomineral
Abbreviations: AFM, atomic-force microscopy; EPM, extracellular polymeric materials; EPXMA, electron-probe X-ray microanalysis; PIXE, proton-induced X-ray emission; TEM, transmission electron microscopy; XRD, X-ray powder diffraction analysis
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INTRODUCTION |
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An alternative mechanism is the coupling of a growth-decoupled, single enzymic step to metal biocrystallization. A Citrobacter sp., originally isolated from metal-polluted soil, overproduces PhoN acid phosphatase (Jeong et al., 1997 , 1998
; Basnakova et al., 1998a
), which is also expressed by several other enterobacteria (Groisman et al., 1992
; Thaller et al., 1995
), and which can mediate metal uptake via enzymically liberated
to precipitate with heavy metals as cell-bound, polycrystalline metal phosphate (Jeong et al., 1997
; Basnakova et al., 1998a
, b
). Previous studies using electron microscopy with immunogold labelling of Citrobacter sp. phosphatase suggested involvement of the phospholipid outer and inner membrane bilayers in the formation of metal phosphate nucleation foci in juxtaposition to the periplasmically localized enzyme (Jeong et al., 1997
). However, extensive mineral deposits were not seen in the periplasmic space but were visible mainly outside the cell body at high metal loadings (Jeong et al., 1997
).
The present study aimed to establish the role of additional exocellular nucleation sites in metal bioprecipitation and crystal growth. A working hypothesis for the process of metal accumulation is formulated, describing this in terms of the localization of the phosphatase and the biochemical and physico-chemical features of the cellular micro-environment. A possible role for phosphatase and cell-surface components in the maintenance of cellular homeostasis is also proposed.
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METHODS |
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Uranium uptake by resuspended cells and examination of metal-loaded cells.
Cells harvested in the mid-exponential phase (2040 ml) were washed twice in isotonic saline (8·5 g NaCl l-1) and resuspended (OD600 0·30·4) in 2 mM trisodium citrate/citric acid buffer, pH 6·9, 5 mM glycerol 2-phosphate, with uranyl nitrate to 1 mM (30 °C). Timed samples (1·25 ml) were centrifuged (22 °C, 5 min, 13700 g) and the supernatant assayed for residual uranyl ion using arsenazo III (Yong & Macaskie, 1995 ). Uranium uptake was calculated as a percentage of bacterial dry weight [mg U (mg bacterial dry weight)-1x100] using a calibration of 0·495 mg dry weight ml-1 for 1 unit of OD600 (Yong, 1996
). The U-loaded cell pellets (each equivalent to 5 ml of culture) were washed twice in isotonic saline (8·5 g NaCl l-1) and once in water, resuspended in 0·20·4 ml water, mixed and placed on a Formvar-coated electron microscope grid, air-dried overnight and examined with a JEOL 1200 EX11 electron microscope (see below), without staining. For monitoring intracellular uranium, U-loaded cells (equivalent to 5 ml of culture) were washed as above, fixed and embedded in LR White as described previously (Jeong et al., 1997
). Sections (80100 nm) cut with a ReichertJung Knifemaker and a microtome (Ultracut E, ReichertJung) were examined without further staining. Specimens were examined using a JEOL 1200 EX11 transmission electron microscope (accelerating voltage 80 kV). Elemental distribution was determined by electron-probe X-ray microanalysis (EPXMA; previously called energy-dispersive X-ray analysis, EDAX) on specimen micro areas (approx. 0·1x0·1 µm) using a JEOL 100 CXII electron microscope (accelerating voltage 100 kV) fitted with a high-resolution scanning attachment, LaB6 filament, 30 mm2 Si(Li) ATW detector and Oxford Instruments Link ISIS microanalysis system as described by Basnakova et al. (1998b
).
To obtain complementary information, whole cells were also examined by atomic-force microscopy (AFM) at BNFL, Preston, UK, using air-dried (2025 °C) mounts on glass slides of metal-loaded cells or cells not challenged with uranyl ion (controls). Specimens were examined using a NanoScope III atomic-force microscope (Digital Instruments, USA). Imaging was carried out using microfabricated Si3N4 tips (nominal spring constant 0·06 N m-1, tip radius approx. 40 nm) in contact mode with the interaction force minimized, as determined by reference to the forcedistance curve as recommended by the manufacturer. Previous studies of biofilms using the AFM (Goddard et al., 1996 ) have shown that some dehydration of the bacterial cells occurs upon air-drying but the technique has potential for imaging of samples of biological origin without pre-treatment (Surman et al., 1996
).
Solid-state methods of biomass examination.
For confirmation of the identity of the accumulated metal phosphate, cells were also examined using proton-induced X-ray emission (PIXE) for elemental mapping of samples and high-sensitivity estimation of elemental content. This bulk technique gives an elemental ratio of the population as a whole, analysing the whole pellet following metal exposure. Cell pellets were washed in water, dried, ground to homogeneity, wet with acetone, then placed on a thin pioloform film on an aluminium target, and dried at room temperature. Quantitative elemental analyses were done using the Oxford Scanning Proton Microprobe (Johansson & Campbell, 1988 ; Watt & Grime, 1989
; Grime & Watt, 1990
; Grime et al., 1991
; Breese et al., 1992
) in the Department of Nuclear Physics, University of Oxford, UK. Elemental maps were obtained of specimen areas of approximately 12 mmx12 mm held within the proton beam (3·0 MeV protons produced using a particle accelerator constructed in the Department of Nuclear Physics, University of Oxford). Matrix major element composition and thickness, which are needed to calculate PIXE corrections, were determined by simultaneously determined Rutherford back-scattering (RBS) spectra. The accuracy of PIXE using the RBS correction was demonstrated by comparison of data obtained by PIXE with that determined by other methods (Tamana et al., 1994
) and sample data were also cross-validated versus EPXMA on common sample fields of U-loaded Citrobacter to check the accuracy of the PIXE technique (Basnakova et al., 1998b
).
The identity of crystalline metal deposits was further confirmed using X-ray powder diffraction analysis (XRD). Metal-loaded biomass samples, ground as above, were examined using a high-precision powder diffractometer in the School of Physics, University of Birmingham (Yong & Macaskie, 1995 , 1998
). Exposure times were up to 16 h to monochromatic Cu K
1 radiation produced using an incident-beam cured-crystal germanium monochromator with asymmetric focusing at 25 °C. The scale error of 2
was 0·007° and the specimen surface displacement was 0·0305 mm, which was checked by a standard reference material (Ag). The powder diffraction patterns were recorded from 5 to 60° (2
) with a step length of 0·05° (2
).
Preparation of extracellular material, and metal uptake by extracted material.
Extracellular polymers were isolated by the method of Morgan et al. (1990) . Cells (usually 1 litre: mid-to late-exponential phase) were harvested by centrifugation and the pellet was washed and resuspended in isotonic saline (200400 ml) and heated at 80 °C (1 h). Cells were removed by centrifugation and cooled supernatant (1 vol.) was treated with 9 vols acetone/ethanol (3:1, v/v; 4 °C, overnight). The white solid was collected under gravity with removal of most of the clear supernatant by aspiration, and finally by centrifugation, washed with acetone and allowed to dry. Before analysis, the samples were washed repeatedly with distilled water, the precipitation step was repeated between each wash and all washings were analysed for inorganic phosphate by a modification of the method of Pierpoint (1957)
as described previously (Jeong et al., 1997
; Yong & Macaskie, 1998
).
Dried sample (several milligrams) was examined using PIXE (as above). Solution 31P NMR spectroscopy (2050 mg sample per tube) was done in a Brüker 400 MHz spectrometer at 161 MHz with a pulse time of 0·91 µs and a pulse recycle delay of 1 s with 85% H3PO4 as the standard and D2O (in a capillary insert) as the field frequency lock. Spectra were acquired before and after metal exposure (1 mM). Initial tests examined uranium binding to the extract but since paramagnetic 238U quenches the NMR signal, tests were also done using 112Cd2+ (1 mM), which is NMR silent. For metal-uptake tests, extracted polymer (20 mg) was placed into a 100 ml conical flask (20 ml) in 20 mM MOPS-NaOH buffer/1 mM citrate buffer (to hold the metal in solution), pH 7·2, and uranyl or cadmium nitrate was added, to 1 mM. The flasks were shaken gently at 30 °C (4 h; time to saturation was determined by prior experiment) and metal-laden material was precipitated and washed as before.
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RESULTS AND DISCUSSION |
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Further studies on the mechanism of exocellular uranyl phosphate accumulation
A technique developed for extraction of EPM (Morgan et al., 1990 ) was used to obtain EPM and other loosely bound, polymeric material as a white precipitate. No further purification was attempted. This material contained a substantial amount of phosphorus, as determined by analysis of repeatedly washed, dried samples using PIXE (3·7 µg P mg-1: Table 1
). The major counterion was Na+, approximately equimolar to the concentration of phosphate. No detectable phosphate was produced in the washings and it was concluded that the extracted material contained bound phosphate species, as confirmed by the 31P NMR spectrum (Fig. 6a
). Most of the phosphate was present as monophosphates (with chemical shifts of 0·3 p.p.m. and 2·0 p.p.m., respectively) with an additional, unassigned peak at 20·5 p.p.m. The 31P spectrum in the region from -5 to 5 p.p.m. was similar to that of the LPS of an E. coli strain described previously (Strain et al., 1983a
, b
), with the 0·3 and 2·0 p.p.m. resonances attributed by these authors to monophosphate groups joined at 1 and 4 positions of the N-acetylglucosamine residues of the lipid A backbone of the LPS (Strain et al., 1983a
, b
). The spectrum (Fig. 6a
) is also very similar to the LPS from Salmonella minnesota strain R345 (Rb) (Batley et al., 1985
), which is believed to produce an almost complete core oligosaccharide (Luderitz et al., 1971
). We conclude that our preparation contained LPS, but since the recovered phosphate (Table 1
) was tenfold less than that for pure LPS (calculated with reference to Klapcinska, 1994
) the preparation was almost certainly not pure. It would have contained extracellular polysaccharidic material, and possibly also outer-membrane phospholipids. However, the latter gave a very broad 31P NMR peak (25 p.p.m. to -25 p.p.m.: Burnell et al., 1980
) in contrast to the sharp peaks in Fig. 6
, while Ferris & Beveridge (1984)
noted that the LPS, containing substantial phosphorus, occurred in outer-membrane vesicles. It should be stressed that the present study did not aim to extract LPS quantitatively; the LPS was co-extracted with other exopolymers. The preparation was not purified further for metal-binding studies using 31P NMR, since the spectrum (Fig. 6a
) indicated few other contaminating phosphate species and a good signal-to-noise ratio was obtained with the crude preparation.
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Metal uptake by the extracted material
Upon addition of to the extract the 31P NMR signal disappeared immediately, corresponding to the disappearance of the phosphate groups from the liquid and into the solid state (Fig. 6b
), and to the immediate appearance of a yellow precipitate at the bottom of the NMR tube. This precipitate, analysed by PIXE, had a composition of (molar ratios) P:Na, 1·1±0·1; U:Na, 14·3±1·1 U: P, 13·0±2·1 (mean±SEM; n=3). Clearly the uranium loading was much greater than stoichiometric and probably represented binding of uranium also to species other than phosphate groups (the LPS was estimated to be only approx. 10% pure: see above). A correspondingly poor XRD powder pattern was obtained from the yellow precipitate (Fig. 6c
) but this was similar to that of Fig. 2
, after data fitting, i.e. it contained hydrogen uranyl phosphate.
It was concluded that the LPS and also other components extracted from Citrobacter N14 accumulate uranyl ion substantially but further tests were not done using because of the quenching effects of the paramagnetic nucleus and because of the precipitation in the experiment. Instead, the diamagnetic (NMR silent) 112Cd2+ was used to monitor metal uptake. In this case the resonances from 2 to -2 p.p.m. disappeared and were replaced by one new resonance, at 3 p.p.m. (Fig. 7
). This is similar to that reported previously by Strain et al. (1983a)
and confirms binding of metal by Citrobacter LPS. Strain et al. (1983a)
noted that the exact peak positions were pH-dependent but a common resonance of 3 p.p.m. for the two phosphate species was seen at pH 6·5. More accurate analysis was not possible in the present case. The extracted material was very viscous when dissolved in the minimum concentration of water (to obtain a good signal-to-noise ratio during examination by NMR); the exact concentration, and the pH, were not determined. Peak positions can be affected by the pH and the Cd:LPS ratio used, as noted by Strain et al. (1983a)
. These authors also noted that, of Mg2+, Ca2+ and Cd2+, the last resulted by far in the largest chemical shift changes, and that the pKa values of LPS phosphate groups in the presence of Cd2+ were shifted to lower pH by approximately 2 units, accounting for the metal-ion-dependent change in the 31P chemical shifts. With the paramagnetic lanthanides (Strain et al., 1983a
) and with Mn2+ (Ferris & Beveridge, 1986
) the peak intensities (but not, substantially, the chemical shifts) decreased with increasing metal concentration. Our attempts to follow 31P chemical shift using paramagnetic
were unsuccessful due to precipitation (above); note that in the previous work (Strain et al., 1983a
; Ferris & Beveridge, 1986
) the metal was added at concentrations several orders of magnitude less than the concentration of LPS phosphorus. The present study was targeted towards high metal loading and the metal was present to excess.
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Requirement for cellular LPS and phosphatase for efficient metal bioprecipitation
The present study implicates LPS as a major site of binding and also of uranyl phosphate nucleation; therefore the localization of the Citrobacter phosphatase was re-examined from a library of electron micrographs obtained in a previous study (Jeong et al., 1997
) to clarify the mechanism of metal uptake. A previous study using Pseudomonas aeruginosa has identified that exocellular membrane vesicles rich in LPS can also serve as a carrier for exocellular alkaline phosphatase; this represents a mechanism for protein export (Kadurugamuwa & Beveridge, 1995
). Fig. 8(a)
shows the localization of a portion of the Citrobacter phosphatase outside the cell, either associated with outer-membrane extrusions or within an indistinct exocellular fuzz. The extracellular material probably acts as a protective immobilizing matrix. Examination of the surface of Citrobacter sp. following uranyl uptake shows some extruded electron-opaque material, but no structural details are apparent (Fig. 8b
). Ferris & Beveridge (1984)
reported binding of metal to outer-membrane vesicles which contained LPS but the extruded material shown in Fig. 8(b)
appears to have little organized structure. It is possible that extracellular vesicles may have been disrupted during sample preparation. Fixation of the cells in a polyacrylamide gel matrix and cryofixation, followed by cryo-ultramicrometry, clearly showed the presence of bacterial cells and extracellular granules of metal phosphate which were, in some cases, associated with the bacterial cells (Basnakova et al., 1998b
).
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The localization of the enzyme is important. Phosphatases are often held within the periplasmic space (Neu & Heppel, 1965 ; Nossal & Heppel, 1966
), and immunogold labelling confirmed this (Jeong et al., 1997
). The extracellular production (export) of enzymes in Gram-negative bacteria is still not completely understood, but the involvement of a secretion-coupled biosynthesis pathway has been suggested with respect to alkaline phosphatase (e.g. Nesmeyanova et al., 1994
) and a secretion process via membrane vesicle production was postulated by Kadurugamuwa & Beveridge (1995)
. In addition to proteolytic modification of the mature subunits in the periplasm, an export mechanism for overproduced enzyme via outer-membrane vesicles was proposed, as visualized in E. coli (Nesmeyanova et al., 1991
) and P. aeruginosa (Kadurugamuwa & Beveridge, 1995
) by electron microscopy and immunogold labelling. The phosphatase activities of the overproducing E. coli (via cloning and overexpression of phoA: Nesmeyanova et al., 1991
, 1994
) and the naturally PhoN-overproducing Citrobacter sp. (this study) were similar. We found no direct evidence for outer membrane vesicle production per se, but possible outer membrane extrusions were visible in some cells in association with phosphatase (Fig. 8a
). The extracted material for the 31P NMR study was washed in acetone but stringent precautions were not taken to exclude membrane phospholipids. However the 31P NMR spectrum of Fig. 6(a)
shows mainly well-defined peaks assigned to monophosphate components of LPS and without the broad peaks associated with 31P NMR spectra of membrane preparations (Burnell et al., 1980
). In the present case, as with previous studies (see above) there is evidence for the production of phosphatase associated with a cell surface matrix of LPS. The role of exopolymeric material as an immobilizing agent for exoenzymes was reported previously (Frolund et al., 1995
). A model can be developed in which the interplay of cellular and microenvironmental factors is crucial to metal biocrystallization. Uranyl ion per se is toxic to the phosphatase and occurrence of the enzyme in a protected environment is likely. The native enzyme is produced as a high-molecular-mass complex, as concluded from unsuccessful attempts to fractionate it using gel-filtration chromatography (Kier et al., 1977
; Jeong et al., 1998
), and non-migration in non-denaturing polyacrylamide gels (Jeong et al., 1998
).
The model for metal biocrystallization by whole cells assumes that the initial event is the formation of a complex between the incoming metal and the monophosphate groups of the LPS. These intercept the metal and thus protect the nearby enzyme for long enough to achieve substrate cleavage and diffusion outward of liberated . Efficient precipitation of uranyl phosphate as the sodium, and not the protonated, form is promoted by capturing the sodium associated with the polymeric material and which was also provided as the counterion with the glycerol 2-phosphate substrate. The initial complexation also forms metal phosphate nucleation sites which are further consolidated by the co-deposition of more incoming metal with the outgoing phosphate, and formation of a polycrystalline material. Metal continues to diffuse inward and phosphate outward, both along a downhill concentration gradient of free ions, since the precipitated metal phosphate is removed from the equilibrium. If the bound enzyme is inhibited, or incoming metal fails to be trapped, a second line of interception can be invoked by the phospholipid groups of the membrane bilayer surrounding the cells and the adjacent reservoir of periplasmic and outer-membrane-bound phosphatase (Jeong et al., 1997
). Hence, there are two pools of both enzyme and nucleation foci, and a dual system for biocrystallization. Two similar, but distinct, forms of the phosphatase were observed previously (Jeong et al., 1998
). It is possible that these represent periplasmic and exocellular forms but confirmation of this awaits further study.
These concepts can explain the fulfilment of two fundamental requirements for future applications to metal waste decontamination. First, very high metal loads can be achieved without fouling by the accumulated precipitate. The architectural arrangement of LPS in native cells is difficult to study, but a meshwork of fibrils or vesicles may hold the metal phosphate crystals in an open structure that allows continued substrate access to the enzyme. Second, the presence of available phosphate groups within the LPS could provide a localized buffering function initially, supplemented by additional phosphate provided by the enzyme. The localized pH could be held reasonably constant irrespective of the pH of the bulk solution. These effects were illustrated by the ability of immobilized cells to liberate phosphate into, and remove uranyl ion from, a solution of acid mine wastewater of pH 3·5, at which pH the phosphatase activity is normally negligible (Macaskie et al., 1997 ). The low pH of (for example) acid mine drainage waters may not, therefore, prove to be too problematic.
This study was originally conceived to develop a mechanistic model to describe metal uptake in biochemical and chemical terms in order to refine the mathematical descriptions (Macaskie et al., 1995 , 1997
) which enable prediction of how the biocatalyst would perform in operation. However, the present findings also allow us to develop a concept of the cell surface outside the outer membrane as a functional physiological compartment. In this model, phosphatase is not exported randomly but is held in association either with extracellular membrane vesicles (Nesmeyanova et al., 1991
; Kadurugamuwa & Beveridge, 1995
) or with strands of LPS. In contrast to PhoA (alkaline phosphatase), which is under the control of the pho regulon and is associated with the supply of phosphate to the cells (Torriani, 1990
), the role of PhoN (acid phosphatase) still remains unclear. It is upregulated by carbon (and also phosphate) starvation (Kasahara et al., 1991
), by shift into anaerobiosis and by osmotic stress (Hallett et al., 1991
), and is regulated via the phoP/phoQ (sensorregulator) regulon (Groisman et al., 1989
; Miller et al., 1989
; Kasahara et al., 1991
, 1992
; Hohmann & Miller, 1994
). Stress could include low pH (Hohmann & Miller, 1994
). A role of the pH 2·5 acid phosphatase was proposed in the hydrolysis of polyphosphate to generate phosphate as a buffer within the periplasm (Dassa et al., 1982
) and a similar pH-homeostatic role could be envisaged for the PhoN phosphatase exocellularly, particularly since its pH optimum is 7·0 and below (Jeong et al., 1998
), and phosphatase scavenges orthophosphate from membrane phospholipids as it is released (Kadurugamuwa & Beveridge, 1995
). The concept of LPS and membrane vesicles as a functional unit in this sense may repay further study but the evidence points to a role for PhoN in the generation of phosphate buffer in the exocellular micro-environment, with metal biocrystallization as a useful side-reaction of this activity.
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ACKNOWLEDGEMENTS |
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Received 27 July 1999;
revised 31 March 2000;
accepted 14 April 2000.