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J Bacteriol, January 1998, p. 159-162, Vol. 180, No. 1
0021-9193/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Dynamics of Iron Uptake and
Fe3O4 Biomineralization during Aerobic and
Microaerobic Growth of Magnetospirillum gryphiswaldense
Dirk
Schüler
and
Edmund
Baeuerlein*
Abteilung Membranbiochemie,
Max-Planck-Institut für Biochemie, D-82152 Martinsried, Germany
Received 27 May 1997/Accepted 22 October 1997
 |
ABSTRACT |
Iron uptake and magnetite (Fe3O4) crystal
formation could be studied in the microaerophilic magnetic bacterium
Magnetospirillum gryphiswaldense by using a radioactive
tracer method for iron transport and a differential light-scattering
technique for magnetism. Magnetite formation occurred only in a narrow
range of low oxygen concentration, i.e., 2 to 7 µM O2 at
30°C. Magnetic cells stored up to 2% iron as magnetite crystals in
intracytoplasmic vesicles. This extraordinary uptake of iron was
coupled tightly to the biomineralization of up to 60 magnetite crystals
with diameters of 42 to 45 nm.
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TEXT |
The most intriguing feature of
magnetic bacteria is the presence of intracellular magnetic inclusions
termed magnetosomes, which are membrane-enveloped crystals of a
magnetic iron mineral (2, 3, 9). Most magnetic bacteria
produce single crystals of the magnetic iron mineral
(Fe3O4), which are aligned in chains, thereby
exerting a magnetic dipole moment to the bacteria. The biological
significance of cellular magnetism is not totally understood, but it is
currently thought that magnetosomes contribute to cellular navigation
by interaction with the Earth's magnetic field ("magnetotaxis") (8). Other physiological functions for magnetite
biomineralization have also been discussed (5, 10, 13).
Intracellular magnetite biomineralization involves the uptake and
accumulation of tremendous amounts of iron. However, knowledge about
iron metabolism in these organisms is limited, and the first reports on
the mechanism of iron transport are contradictory (14, 15).
Also, the physiological relationship between iron uptake and its
subsequent storage in the form of magnetite crystals, species specific
in form and size, remains obscure. When magnetite is formed during
growth, whether the internalized iron is directly converted to
magnetite, or if iron can be accumulated and stored before in a
nonmagnetic form are open questions.
In the present study, we tried to find the precise physiological
conditions under which magnetite biomineralization occurs in the
magnetic bacterium Magnetospirillum gryphiswaldense MSR-1 (16, 19). Cells of M. gryphiswaldense (DSM 6361)
are able to form up to 60 cubo-octahedral crystals of magnetite. The
individual particles are enveloped by a membrane with a distinct
protein and lipid profile (1, 17). In a previous study, a
high potential for the transport of ferric iron into iron-starved cells
was detected in this bacterium (18). Energy-dependent iron
uptake obeyed Michaelis-Menten-kinetics but did not involve
siderophore-like compounds. In the present study, we characterized the
dynamics of magnetite formation during growth using a light-scattering method for the quantification of magnetism. We also present data demonstrating that iron uptake in M. gryphiswaldense is
tightly coupled to the induction of magnetite biomineralization.
Dynamics of magnetite formation during aerobic and microaerobic
growth.
In initial growth experiments, cells were agitated at
30°C in free gas exchange with air in a growth medium supplemented
with 30 µM ferric citrate, as previously described (18).
Cell growth was determined by measuring the optical density at 400 nm.
The average magnetic orientation of cell suspensions ("magnetism") was assayed by an optical method as previously described
(20). In this method, cells are aligned at different angles
relative to the light beam by means of an external magnetic field. The ratio of the resulting scattering intensities
(Cmag) correlated well with the average number
of magnetic particles and could be used for the semiquantitative
evaluation of magnetite formation (for practical purposes,
Cmag = 0 was assumed for nonmagnetic cells;
Cmag = 1 then corresponded to approximately 10 particles per cell). At moderate agitation (120 rpm), the average
number of magnetic particles in the cells varied during the growth
period (Fig. 1). During early growth, the
average cellular magnetism gradually declined. This was likely due to
the dilution of magnetic particles in the culture by cell division
while the formation of new magnetite crystals was inhibited. After an
incubation time of about 25 h, only weak residual magnetism was
detected in the culture. This transient decrease in magnetosome content
was followed by a rapid increase in magnetite formation when growth
proceeded exponentially.
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FIG. 1.
Variations of magnetism in a culture of M. gryphiswaldense during growth monitored by changes in differential
light scattering (Cmag). Magnetic cells were
inoculated into a medium which was constantly agitated and contained 30 µM ferric citrate.  , Magnetism (M);  , optical density (OD).
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To study the effect of oxygen on magnetite biomineralization, cells
were grown in a 5-liter laboratory fermentor vessel (Bioflow
III; New
Brunswick Scientific) for determining continuously the
concentration of
dissolved oxygen. In these experiments, nonmagnetic
cells which had
been grown in the absence of an added iron source
were used as an
inoculum (initial cell concentration, 5.0 × 10
6 cells
ml
1) in a growth medium containing 30 µM ferric
citrate. At an aeration
rate of 0.5 liters of air/min, the oxygen level
in the culture
was high during the initial growth period (aerobic
growth). As
growth proceeded, the medium became gradually depleted of
oxygen
(Fig.
2A). Cells were nonmagnetic
during aerobic growth but started
to produce
Fe
3O
4 immediately after microaerobic conditions
were
attained, with an oxygen concentration of about 1 to 3%
saturation
(2 to 7 µM O
2 at 30°C). At an increased
aeration rate of 2 liters
of air/min, aerobic growth was prolonged,
resulting in a higher
cell yield but a reduced magnetite content in
cells (Fig.
2B).
At an aeration rate of 3.0 liters of air/min, however,
the oxygen
concentration did not decrease below 5% saturation. Cells
eventually
grew after a prolonged lag phase from a large inoculum
(initial
cell concentration, 10
7 cells ml
1),
but magnetic cells were not detected at any stage of growth.
Aeration
rates >3 liters of air/min totally inhibited growth.
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FIG. 2.
Dissolved oxygen concentration (O2) ( ),
cell density (OD) ( ), and magnetism (M) () in cultures of
M. gryphiswaldense during growth at different aeration
rates. Aeration was kept constant throughout each experiment: 0.5 liter
of air/min (A) and 2.0 liters of air/min (B). The experiments were
started by the inoculation of nonmagnetic cells into the growth
medium.
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Relationship between iron uptake and magnetite formation.
Growth, magnetism, and iron content of cells were measured
simultaneously throughout growth. The iron content of cells was determined by using the radioactive isotope 55Fe in a
manner similar to that described previously (18). Briefly, at various time intervals, 1-ml samples were withdrawn, added to 5 ml
of 0.1 M LiCl-5 mM EDTA, and filtered (0.45-µm-diameter pore size;
Sartorius). The filters were washed once with 5 ml of 0.1 M LiCl-5 mM
EDTA and then dried. Radioactivity on the filters was determined using
a liquid scintillation counter (scintillant, Ultima Gold; Packard
Instrument). The iron content was calculated from the specific
radioactivity supplied in the medium. A culture density of 1 at 400 nm
corresponded to 0.21 mg (dry weight)/ml. Nonmagnetic cells were
inoculated into two identical batch cultures containing 100 ml of
medium without added ferric citrate (initial cell concentration,
5.0 × 106 cells ml1). Cultures were
incubated in loosely capped flasks at 30°C with moderate agitation
(120 rpm). In experiment A, (Fig. 3A)
radioactive 55FeCl3 (DuPont-NEN) was added
immediately after inoculation to a final concentration of 30 µM
(specific activity, 20 MBq mg1). In a previous experiment
(18) it was shown by the Ferrozine assay that iron, supplied
to the culture as Fe(II)SO4 was predominantly in the ferric
form but remained soluble throughout the experiment.
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FIG. 3.
Cell density (OD) (), cellular magnetism (M) (),
and intracellular iron content (Fe) ( ) of M. gryphiswaldense during growth. The experiment was started by
simultaneous inoculation of nonmagnetic cells for both panels. Iron was
added to 30 µM as 55FeCl3 (in 1 N HCl) at the
time of inoculation (A) and after 14.5 h (B) as indicated by
arrows. The figures given do not accurately reflect the iron content of
the cells since the iron content of the nonmagnetic inoculum was not
taken into account.
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Uptake of iron did not occur continuously during growth but was closely
coupled to the initiation of magnetite formation (Fig.
3A). A transient
peak in iron content during initial growth was
followed by a decrease
in the cellular iron content. During further
growth, the iron content
of the nonmagnetic cells remained low
at 0.02 to 0.06% (dry weight)
(
11), indicating that iron uptake
and growth were balanced
while cells did not produce magnetite.
Cellular magnetism was detected
after 14 h, indicating that microaerobic
conditions were reached,
as seen before in Fig.
2. The initiation
of magnetite formation was
accompanied by a drastic increase in
iron uptake.
In experiment B (Fig.
3B), first the radioactive iron was omitted from
the medium, which then still contained about 1 µM Fe.
This iron
concentration was sufficient for growth but prevented
significant
magnetite production by cells (
18).
55FeCl
3 (30 µM) was added here at the time
when the cells began
microaerobic growth, as indicated by detectable
magnetism in experiment
A (Fig.
3A). The cells started to take up the
radioactive iron
immediately after its addition (Fig.
3B).
Concomitantly, the cellular
magnetism rapidly increased, indicating
that the ingested iron
was converted to magnetite without apparent
delay. About 5 to
10 min after the addition of iron, magnetite
formation could be
detected by a change of the light scattering of the
culture in
a magnetic field (
20). At the end of growth,
cells from experiment
B (Fig.
3B) had a much higher iron content and
were more magnetic
than cells from experiment A (Fig.
3A), in which the
iron concentration
was kept constantly high at 30 µM. For electron
microscopy, samples
of cells were taken from the culture and the cells
were killed
by the addition of formaldehyde (1 drop to 1 ml of cell
suspension).
Samples were concentrated and viewed with a Philips CM 10 transmission
electron microscope at 100 kV. Shortly after the addition
of iron
in experiment B (Fig.
3B), the cells contained numerous tiny
intracellular
electron-dense particles arranged in a loose chain (Fig.
4A).
These crystallites were imperfect in
morphology and predominantly
in the superparamagnetic size range (5 to
20 nm) (
7), while
mature crystals usually had a perfect
cubo-octahedral shape and
were 45 nm in diameter (Fig.
4B).
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FIG. 4.
Electron micrographs of magnetosomes in cells of
M. gryphiswaldense. (A) Chain of magnetosomes from a cell
grown in the presence of a steady concentration of 30 µM
FeCl3. Mature crystals, which are prevalently located in
the middle of the chain, are cubo-octahedral and 45 nm in diameter. (B)
Crystals present in cells about 30 min after the induction of
Fe3O4 biomineralization by the addition of 30 µM FeCl3. The immature particles are 5 to 20 nm in
diameter and fit predominantly into the superparamagnetic size range
(7).
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We described here the physiological conditions in the presence of which
magnetite crystals (Fe
3O
4) are biosynthesized.
Whereas
cells tolerated higher oxygen concentrations for growth,
Fe
3O
4 was produced only during microaerobic
growth. Magnetite formation
was induced in nonmagnetic cells by a low
threshold oxygen concentration
of about 2 to 7 µM O
2
(30°C). The dependence of magnetite formation
on low oxygen levels is
in accordance with earlier findings in
Magnetospirillum
magnetotacticum (
6). At low E
h, the
inorganic
synthesis of magnetite (Fe
3O
4) at
neutral pH is known to be thermodynamically
favored compared to that of
other crystalline iron oxide phases
like Fe
2O
3
(
4). Hence, it appears likely that the development
of
microaerophilic conditions directly affects the physiochemical
conditions in the interior of magnetosome vesicles, favoring the
precipitation of Fe
3O
4. Magnetite formation was
tightly coupled
to a drastic increase in iron uptake, whereas the iron
content
of nonmagnetic cells was very similar to that reported for
other
nonmagnetic bacteria (
11,
12). Both the uptake of iron
and
the formation of magnetite were stimulated by microaerobic
conditions.
Apparently, the iron taken up by the cells was rapidly
converted
to Fe
3O
4 with no delay. Thus, our
results do not support the idea
that the formation of magnetite
crystals is preceded by the accumulation
of a large iron pool stored in
a nonmagnetic form.
Interestingly, iron-depleted cells had a higher potential for iron
accumulation and magnetite formation compared to cells
adapted to high
iron concentrations. This finding suggests the
regulation of iron
uptake. It may be assumed that uptake system(s)
would be derepressed in
iron-deficient cells, resulting in a transiently
increased potential
for iron uptake if iron-rich conditions were
encountered. Since
iron-depleted bacteria were able to form magnetite
crystals and to
internalize iron without lag immediately upon
its addition, we suppose
that any structures and/or enzymes potentially
involved in iron uptake
and magnetite synthesis are preexisting
in the cell and do not require
activation or induction by the
presence of high iron concentrations in
the medium.
These results allow us now to cultivate
M. gryphiswaldense
and to prepare and purify its magnetosomes in large amounts for
studying the proteins and phospholipids of the magnetosome membrane
(unpublished data).
| |
ACKNOWLEDGMENTS |
We thank Dennis A. Bazylinski for critical reading of the
manuscript.
This work was supported by a grant (E.B.) from the Deutsche
Forschungsgemeinschaft (DFG) (Schwerpunktsprogramm "Bioinorganic Chemistry: Transition Metals in Biology and Their Coordination Chemistry").
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Membranebiochemistry, Max-Planck-Institute for Biochemistry, D-82152 Martinsried, Germany. Phone: 49-89-8578 2359. Fax: 49 89-8578 3777. E-mail: baeuerlein{at}biochem.mpg.de.
Present address: Department of Microbiology, Immunology & Preventive Medicine, Iowa State University, Ames, IA 50011.
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REFERENCES |
| 1.
|
Baeuerlein, E., and D. Schüler.
1995.
Biomineralisation: iron transport and magnetite crystal formation in Magnetospirillum gryphiswaldense.
J. Inorg. Biochem.
59:107.
|
| 2.
|
Balkwill, D. L.,
D. Maratea, and R. P. Blakemore.
1980.
Ultrastructure of a magnetotactic spirillum.
J. Bacteriol.
141:1399-1408[Abstract/Free Full Text].
|
| 3.
|
Bazylinski, D. A.
1995.
Structure and function of the bacterial magnetosome.
ASM News
61:337-343.
|
| 4.
|
Bell, P. E.,
A. L. Mills, and J. S. Herman.
1987.
Biogeochemical conditions favoring magnetite formation during anaerobic iron reduction.
Appl. Environ. Microbiol.
53:2610-2616[Abstract/Free Full Text].
|
| 5.
|
Blakemore, R. P., and N. Blakemore.
1991.
Magnetotactic magnetogens., p. 51-68. In
R. B. Frankel, and R. P. Blakemore (ed.), Iron biominerals.
Plenum Press, New York, N.Y.
|
| 6.
|
Blakemore, R. P.,
K. A. Short,
D. A. Bazylinski,
C. Rosenblatt, and R. B. Frankel.
1985.
Microaerobic conditions are required for magnetite formation within Aquaspirillum magnetotacticum.
Geomicrobiol. J.
4:53-71.
|
| 7.
|
Butler, R. F., and S. K. Banerjee.
1975.
Theoretical single-domain grain size range in magnetite and titanomagnetite.
J. Geophys. Res.
80:252-259.
|
| 8.
|
Frankel, R. B., and D. A. Bazylinski.
1994.
Magnetotaxis and magnetic particles in bacteria.
Hyperfine Interact.
90:135-142.
|
| 9.
|
Gorby, Y. A.,
T. J. Beveridge, and R. P. Blakemore.
1988.
Characterization of the bacterial magnetosome membrane.
J. Bacteriol.
170:834-841[Abstract/Free Full Text].
|
| 10.
|
Guerin, W. F., and R. P. Blakemore.
1992.
Redox cycling of iron supports growth and magnetite synthesis by Aquaspirillum magnetotacticum.
Appl. Environ. Microbiol.
58:1002-1109.
|
| 11.
|
Hartmann, A., and V. Braun.
1981.
Iron uptake and iron limited growth of Escherichia coli K-12.
Arch. Microbiol.
130:353-356[Medline].
|
| 12.
|
Lankford, C. E.
1973.
Bacterial assimilation of iron.
Crit. Rev. Microbiol.
2:273-331.
|
| 13.
|
Mann, S.,
N. H. C. Sparks, and R. G. Board.
1990.
Magnetotactic bacteria: microbiology, biomineralization, palaeomagnetism and biotechnology.
Adv. Microbiol. Physiol.
31:125-181[Medline].
|
| 14.
|
Nakamura, C.,
T. Sakaguchi,
S. Kudo,
J. G. Burgess,
K. Sode, and T. Matsunaga.
1993.
Characterization of iron uptake in the magnetic bacterium Aquaspirillum sp. AMB-1.
Appl. Biochem. Biotechnol.
39/40:169-177.
|
| 15.
|
Paoletti, L. C., and R. P. Blakemore.
1986.
Hydroxamate production by Aquaspirillum magnetotacticum.
J. Bacteriol.
167:153-163[Abstract/Free Full Text].
|
| 16.
|
Schleifer, K. H.,
D. Schüler,
S. Spring,
M. Weizenegger,
R. Ammann,
W. Ludwig, and M. Köhler.
1991.
The genus Magnetospirillum gen. nov., description of Magnetospirillum gryphiswaldense sp. nov. and transfer of Aquaspirillum magnetotacticum to Magnetospirillum magnetotacticum comb. nov.
Syst. Appl. Microbiol.
14:379-385.
|
| 17.
|
Schüler, D., and E. Baeuerlein.
1997.
Iron transport and magnetite crystal formation of the magnetic bacterium Magnetospirillum gryphiswaldense.
J. Phys. IV France
7:C1-647-C1-650.
|
| 18.
|
Schüler, D., and E. Baeuerlein.
1997.
Iron-limited growth and kinetics of iron uptake in Magnetospirillum gryphiswaldense.
Arch. Microbiol.
166:301-307.
|
| 19.
|
Schüler, D., and M. Köhler.
1992.
The isolation of a new magnetic spirillum.
Zentralbl. Mikrobiol.
147:150-151.
|
| 20.
|
Schüler, D.,
R. Uhl, and E. Baeuerlein.
1995.
A simple light-scattering method to assay magnetism in Magnetospirillum gryphiswaldense.
FEMS Microbiol. Lett.
132:139-145.
|
J Bacteriol, January 1998, p. 159-162, Vol. 180, No. 1
0021-9193/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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