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Applied and Environmental Microbiology, November 2000, p. 5092-5098, Vol. 66, No. 11
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Heavy Metal Coprecipitation with Hydrozincite
[Zn5(CO3)2(OH)6] from
Mine Waters Caused by Photosynthetic Microorganisms
Francesca
Podda,1
Paola
Zuddas,1
Andrea
Minacci,2
Milva
Pepi,2 and
Franco
Baldi2,*
Department of Earth Sciences, University of
Cagliari, 09127 Cagliari,1 and
Department of Environmental Sciences, University Cà
Foscari, 30122 Venice,2 Italy
Received 24 April 2000/Accepted 21 July 2000
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ABSTRACT |
An iron-poor stream of nearly neutral pH polluted by mine tailings
has been investigated for a natural phenomenon responsible for the
polishing of heavy metals in mine wastewaters. A white mineralized mat,
which was determined to be hydrozincite
[Zn5(CO3)2(OH)6] by
X-ray diffraction analysis, was observed in the stream sediments mainly
in spring. The precipitate shows a total organic matter residue of 10%
dry weight and contains high concentrations of Pb, Cd, Ni, Cu, and
other metals. Scanning electron microscopy analysis suggests that
hydrozincite is mainly of biological origin. Dormant photosynthetic
microorganisms have been retrieved from 1-year-old dry hydrozincite.
The autofluorescent microorganisms were imaged by a scanning confocal
laser microscope. A photosynthetic filamentous bacterium, classified as
Scytonema sp. strain ING-1, was found associated with
microalga Chlorella sp. strain SA1. This microbial
community is responsible for the natural polishing of heavy metals in
the water stream by coprecipitation with hydrozincite.
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TEXT |
Abandoned mines present a high
environmental hazard in all countries today. In several parts of the
world where mining activities have shut down, the problem of the
control and reclamation of polluted areas for new activities arises.
The polishing of metals from a mining area is a difficult task. The
transformation of metals into harmless species or their removal in a
suitable recycled mineral form such as carbonates (1, 15) is
a possible solution for the remediation of a mining area. Therefore,
research in this field continues, with the isolation of new strains
with more-successful mechanisms for the reduction of metal toxicity.
At Ingurtosu (southwestern Sardinia, Italy) lead and zinc sulfide ore
deposits were mined until 1968 and tailings were deposited along the
Rio Naracauli creek. The Montevecchio-Ingurtosu deposit consists of
galena-sphalerite veins in a quartz gangue containing iron, calcium,
and magnesium carbonate minerals. Pyrite, chalcopyrite, barite,
cerussite, and anglesite are the most commonly associated minerals
(22). Previous studies in this area have shown that waters
are highly polluted by heavy metals, in spite of their near-neutral pH
(4-6, 30).
The aim of this work was to study the fate of leached toxic metals from
sulfide ore tailings in the upper part of the Rio Naracauli creek,
where natural polishing of metals in waters occurred as a result of a
very large growth of photosynthetic microbial populations that
colonized sediments in spring and deposited a white mat on the creek
bed. The role of this photosynthetic community adapted to toxic metals
and the mechanism of metal sequestration were studied.
Study area and samplings.
The Rio Naracauli flows in a 30.2 km2 basin west of the Ingurtosu mine in the Arburese mine
district in southwestern Sardinia (Fig.
1). The river is about 8.2 km long and
flows into the western Mediterranean Sea. The Rio Naracauli has a very
limited flow, particularly in the upper part. Upstream it receives
drainage from mine tailings on the left, and downstream it receives
drainage from three adits: the Rio Pitzinurri (outlet A), the Ledoux
mine gallery (outlet B), and the Rio Bau (outlet C). The
hydrogeological details of this area have been reported by Pala et al.
(21).
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FIG. 1.
Schematic map of the sampling area, with tailings
distribution (hatched areas). Samples 1 to 11 ( ) are from the Rio
Naracauli stream; samples A to C were collected in the tributaries
before the inflows.
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A total of 14 stations (stations 1 to 11 in the creek and stations A to
C in the three tributaries) were chosen along 3.4
km of the Rio
Naracauli (Fig.
1). Station 1 was located at the
tailings pond. In
stations 2 to 4, a photosynthetic microbial
population visibly
encrusted the sediments with a green mat in
spring, which developed
into white material, particularly at stations
3 and 4. This is an
annual event that varies in intensity depending
on the meteorological
conditions. The white precipitate is then
mechanically transported away
by rainfalls. Stations 5 to 7 were
located after the Rio Pitzinurri
tributary (sample A). In the
sediments of these stations white
precipitate residues were still
visible. Stations 8 to 10 were located
after the Ledoux gallery
(sample B). Station 11 was located downstream
from the Rio Bau
tributary (sample C), where white deposits were not
observed.
Hydrozincite determination and distribution.
The white
mineralized material at stations 3 and 4 was collected for
mineralogical, chemical, and microscopic analyses in April 1997, concomitantly with a photosynthetic microorganism bloom. At station 4 the precipitated material was very abundant. The mineralogical
characterization of the mineral precipitate was performed by X-ray
powder diffraction (XRD) with a diffractometer (Philips; model PW 1710)
with CuK
radiation. The mineral was determined to be crystalline
hydrozincite
[Zn5(CO3)2(OH)6]
(Fig. 2). Minor minerals, such as quartz
and calcite, were also observed. XRD spectra also showed a significant
background noise, which was due to the presence of organic matter.
Samples of the white solid precipitate were also collected at other
sites along the Rio Naracauli creek, and their XRD spectra showed a
significant decrease in crystalline hydrozincite from sample 6 to
sample 7 (Fig. 2). A white, soft, unconsolidated mud was also collected from stations 8 to 10. This precipitate was identified as amorphous material made up of Zn, Si, and oxygen (30). From sample 8 to the last station, hydrozincite was not detected.
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FIG. 2.
XRD spectra of some selected precipitates in Rio
Naracauli, with relative intensity peaks of hydrozincite (H), calcite
(Cal), and quartz (Qz).
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The concentrations of the main selected chemical components in
hydrozincite precipitates from station 4 are reported in Table
1. The organic carbon content was
determined by the oxidation
method (
3). A white solid sample
(1.0 g) was finely ground
and dried at 105°C. The sample was treated
with a mixture of 20
ml of K
2Cr
2O
7
(0.17 M) and 20 ml of concentrated H
2SO
4 at
room
temperature. Excess potassium dichromate was titrated with ferrous
sulfate according to the Walkley-Black method (
28).
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TABLE 1.
Organic mattera metals, and
sulfate concentrations in the hydrozincite sediment sampled at station
4 in Rio Naracauli
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Elemental characterization of the white precipitate materials was
performed with 0.25 g of the sample. A high-purity mixture
of 5 ml
of concentrated HCl and 7.5 ml of concentrated HNO
3 was
added to the solid sample in a Teflon vessel. The sample was then
heated by microwave (model CEM-MDS 2100) for 25 min. The sample
was
then cooled, 2 ml of H
2O
2 (30% by volume) was
added, and the
sample was once more microwaved for 17 min for final
digestion.
The final solution was filtered and made up to 25 ml using
Milli
Q water. All the elements were determined by inductively coupled
plasma-atomic emission spectroscopy. As shown in Table
1 the
Zn content
in the analyzed sediment, considering the presence
of contaminant
phases, is in good agreement with the theoretical
formulae of
hydrozincite (
17). Hydrozincite coprecipitates high
concentrations of other toxic elements such as Pb, Cd, Cu, and
Ni, in
agreement with the decrease in metals observed in the stream
waters.
Coprecipitation of these metals occurred in concomitance
with the
formation of hydrozincite, by a mechanism similar to
that previously
reported for strontium precipitation in calcite
mineral
(
12). The high Ca concentration was due to the presence
of
calcite and secondary CaSO
4 traces in hydrozincite. The
total
Fe concentration was very low, as observed in the depositing
waters.
Microbiological evidence of hydrozincite formation.
Further
mineralogical analyses were performed on a Cambridge 250 MK3 scanning
electron microscope (SEM) associated with an energy-dispersive X-ray
system (EDX; LINK AN 10/55S). The biological origin of hydrozincite was
clearly pointed out by SEM observations (Fig.
3). The precipitate of minerals around
biological structures formed a network of microscopic tubing (Fig. 3A).
In detail the mineral seems to encrust filamentous bacteria (Fig. 3B).
Broken filaments showed an open internal diameter of 6.0 ± 0.8 µm. At a greater magnification, SEM analysis showed a small amount of "naked" sheath under the inorganic encrustation (Fig. 3C, left). In
more detail, spherical and rough particles of different sizes strongly
adhered to the bacterial sheath (Fig. 3C, right). Hydrozincite microspherical particles are approximately 0.2 µm in diameter. Spherical particles are cemented together in long tubing and produce a
very hard crust, coating the pebbles of the bottom sediments. EDX
element analysis of the precipitate confirmed the presence of Zn, with
minor Si, Ca, and S in the mineral phase (Fig. 3D).
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FIG. 3.
Photos of white precipitate obtained by SEM. (A)
Formation of tubing network of hydrozincite produced by microorganisms.
Bar, 400 µm. (B) Detail of a tubing section giving a better view of
the biological origin of hydrozincite. Bar, 4 µm. (C) (Left) Further
details with naked parts of a sheath. Bar, 10 µm. (Right) Close-up
detail of the sheath on a framed rectangle. (D) EDX elemental analysis
of a sheath encrustation revealing high Zn concentrations.
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To further demonstrate the biological origin of hydrozincite in Rio
Naracauli, about 2.0 g of the 1-year-old solid sample,
stored in
dark, dry conditions and sterile vials, was incubated
in a flask under
dim light at room temperature, by adding a 1:10-diluted,
conventional
BG-11 medium (American Type Culture Collection medium
616) for
cyanobacterial isolation. After a few days of incubation,
filaments
formed from the white precipitate (Fig.
4A) and were
coated by green filaments
(Fig.
4B).
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FIG. 4.
(A) Photograph of a piece of old, dried, consolidated
hydrozincite with dark spots, with cyanobacterium Scytonema
sp. in a dormant state. Bar, 0.5 cm. (B) The same piece of hydrozincite
after a period of incubation in BG-11 medium (diluted 1:10) under dim
light, clearly showing growth of Scytonema sp., which was
entrapped in the hydrozincite matrix and which forms blue-green
filaments. Bar, 0.5 cm. (C) In the transmission mode the
Scytonema sp. clearly grows out of the hydrozincite tubing.
A thick-wall heterocyst (short arrow) is clearly visible. Bar, 12, µm. (D) The same image by SCLM shows single autofluorescent cells of
a Scytonema sp. Autofluorescence is due to chlorophyll
a encapsulated in the sheath. Where the heterocyst occurs
(short arrow), no autofluorescence appears, since the photosynthetic
apparatus is degenerated. Moreover, hydrozincite emits fluorescence
(long arrow). Bar, 12 µm. (F) Coculture of Scytonema sp.
and the microalga Chlorella sp. The cyanobacterium shows
filaments of different diameters. Bar, 12 µm. (G) The same image
taken by SCLM (the sheath of the Scytonema sp. becomes empty
and wide with aging and autofluorescence [arrow]). Bar, 12 µm.
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The autofluorescence of the microbial community due to photosynthetic
pigments was observed by confocal microscopy. The white
solid specimen
was soaked in sterile buffered water and then observed
without any
treatment under light transmission and in fluorescence
modes with
scanning confocal laser microscopy (SCLM) (Bio-Rad
Microscience
Division; model MRC-500); the microscope was equipped
with a
krypton-argon laser, with maximum emission at 488 nm. The
cells emit
red autofluorescence due to the photosynthetic pigments
contained in
them. Three-dimensional images were taken with the
procedure reported
by Baldi et al. (
2). Images were collected
in the
transmission mode (Fig.
4C and F) and in the fluorescence
mode (Fig.
4D
and G). A terminal heterocyst due to the lack of
photosynthetic
apparatus (Fig.
4D) was present, as confirmed by
the presence of a
thick-wall cell observed under the light transmission
mode (Fig.
4C).
From these two photographs it was evident that
a new filament of
cyanobacterium was produced inside the tubing
cavity, which was formed
by hydrozincite precipitation around
the microbial
sheath.
The cyanobacterium was identified as
Scytonema sp. strain
ING-1 based on its morphological characteristics. It forms uniseriated
and sheathed trichomes with false branches made up of hormogonia
and a
single terminal heterocyst. It colonizes mainly freshwater
benthic
habitats (
7). In flask cultures,
Scytonema sp.
strain
ING-1 produced gel-holding masses of trichomes strongly adhering
to glass surfaces and/or sediment gravels of the Rio Naracauli
creek.
After 7 days of incubation another photosynthetic coccoid
microorganism, with a diameter of 4.0 to 4.5 µm was observed (Fig.
4F
and G). The microalga was classified as
Chlorella sp. strain
SA-1 by the size and spherical shape of a nonmotile cell with
a
peripheral chloroplast. The microalga remained as a single cell,
well
anchored to the cyanobacterium sheath network, until both
microorganisms were isolated under laboratory
conditions.
Pigment analysis was carried out on the following three microorganisms:
Scytonema sp. strain ING-1,
Chlorella sp. strain
SA-1
grown in a liquid BG-11 medium, and the reference strain
Chlamydomonas reinhardtii, grown in R medium
(
19). Each culture was filtered
through nitrocellulose
filters (0.2-µm pore size; Sartorius).
Cells were harvested, and
pigments were extracted by a 3-ml acetone-methanol
(7:2) solution.
Absorption spectra of pigments were recorded with
a spectrophotometer
from 300 to 800 nm (Shimadzu; UV-360). In
the cyanobacterium, pigment
spectra showed peaks of phycocyanin
and phycoerythrocyanin at 617 and
582 nm, respectively, plus a
peak of chlorophyll
a at 663 nm. Large amounts of carotenoids
were also produced, as confirmed by
absorption peaks at 432 and
478
nm.
In the coccoid
Chlorella sp. the pigment emits at 662 nm
corresponding to chlorophyll
a, and a large peak at 439 nm,
due to
carotenoids, was also determined. This pigment spectrum was
similar
to that of
C. reinhardtii, confirming that the
coccoid photosynthetic
cell was a
microalga.
The Alcian blue stain technique (
20) was used to determine
the presence of acidic polysaccharide produced by the cultivated
photosynthetic community isolated from the white precipitates
in a
BG-11 medium, while carbofuchsin was used for counterstaining.
The
specimen was heat fixed on a glass slide, and the blue-stained
specimen
was observed under a light transmission microscope. Under
the
fluorescence mode, the precipitate mineral coating the sheath
was also
autofluorescent (Fig.
4D), as shown by the emission of
a strong green
light. This observation suggested the presence
of a molecular secretion
by the microorganism involved in the
formation of hydrozincite. In fact
after several weeks of incubation
of the
Scytonema sp.,
older sheaths became progressively empty
and wide and emitted a similar
green autofluorescence (Fig.
4G).
The polysaccharide sheath also became
acidic, as confirmed by
staining with Alcian blue. Hormogonia and young
narrow filaments
were not stained by Alcian blue. In the formation of
hydrozincite,
the role of
Chlorella sp. is probably that of
reinforcing the
filament structures and sequestering even more cations,
because
of its own acidic polysaccharide capsule, which is also
positive
to Alcian blue
staining.
Water analysis.
During the same sampling campaign, 14 superficial water samples from the Rio Naracauli were collected at the
11 stations plus the three outflows (A, B, and C). The water samples
were filtered through 0.45-µm-pore-size polycarbonate filters by a
metal-free filtering apparatus (Nucleopore). Filtered water samples
were collected into previously acid-cleaned polyethylene bottles. For metal analysis, water subsamples were acidified by HNO3
(Aristar grade; Merck) to reach a final 1% concentration. For anion
analyses, nonacidified water subsamples were stored in the dark at
4°C.
Parameters such as the temperature (degrees Celsius), conductivity
(microsiemens per centimeter), pH, redox potential (millivolts),
and
alkalinity of the waters were measured in situ (
14). The
major cations, Ca
2+, Mg
2+, Na
+,
K
+, Zn
2+, and Cd
2+, were determined
by inductively coupled plasma-atomic emission
spectroscopy (ARL3520).
Other trace elements, Pb
2+, Cu
2+,
Ni
2+, and Co
2+, were determined by inductively
coupled plasma-mass spectroscopy
(Perkin-Elmer; ELAN 5000). These
analytical procedures and their
reproducibilities and sensitivities
have been reported by Cidu
(
8).
In filtered nonacidified subsamples, the major anions (CI
and SO
42) were analyzed by high-pressure
liquid chromatography on an instrument
(Dionex) equipped with an
AS12A-Sc column, a conductivity detector
(model CDM1), and a
carbonate-bicarbonate mobile phase (2.7 and
0.3 mM) at 1.5 ml
min
1 (
16). Analytical data for the waters from
stations 1 to 8,
sampled in the area affected by hydrozincite
deposition, are reported
in Table
2.
Based on the major components, the waters show a dominant
CaSO
4-MgSO
4 chemical composition, except for
sample A, which represents
the Rio Pitzinurri tributary. At stations 1 and 2 the main cation
in the water was Zn
2+; in the other
stations the main cations were Ca
2+ and Mg
2+.
Along the analyzed stream section, the salinity as total dissolved
solids (TDS) shows large variations. Between stations 1
and 2, the TDS
decrease was due to the dilution effect of sulfate
and metal-poorer
waters. Conversely, TDS increased between stations
2 and 3 due to a
significant input of HCO
3-rich waters from
tailings located along the stream (Fig.
1).
A rapid increase in the pH
took place from station 2 to stations
3 and 4 due to photosynthetic
activity. This process was associated
with a decrease in
HCO
3, especially between stations 3 and 4, where hydrozincite precipitation
occurred. Toxic metals
Zn
2+, Cd
2+, Pb
2+, Ni
2+,
and Cu
2+ decreased dramatically in waters at stations 3 and
4, concurrent
with carbonate formation, and more gradually at the other
stations.
From the first station to the fourth Zn, Cd, Pb, and Cu
contents
in waters decreased by 92, 87, 94, and 84%
respectively.
Chemical speciation, partial CO
2 pressure
(pCO
2), and equilibrium calculations for water samples were
performed by the WATEQP
computer program (C. A. J. Appelo,
WATEQP
a computer program for
equilibrium calculations of water
analysis, Institute of Earth
Sciences, Free University of Amsterdam,
1988). The saturation
index (SI) with respect to the mineral phases was
calculated as
log (IAP/
Ks), where IAP is the
ionic activity product and
Ks is
the equilibrium
constant at the water temperature. The SI in the
waters was also
calculated for the following carbonates: hydrozincite,
calcite,
smithsonite (ZnCO
3), and cerussite (PbCO
3).
Hydrozincite
saturation equilibrium was reached at station 3 (SI
H =
0.04),
and it was due to a significant input
of HCO
3-rich waters from other tailings
between stations 2 and 3 along
the Rio Naracauli. The further
oversaturation equilibrium conditions
(SI
H = 1.50)
reached in station 4 are not due to chemical inputs
or outputs but to
the photosynthetic activities of identified
microorganisms. In the
waters of stations 3 and 4, chemical variations
were observed only for
HCO
3, Zn
2+, Pb
2+, and
Cd
2+, while other ions did not change. The SI with respect
to hydrozincite
formation was also achieved in stations 6 and 7. Calcite equilibrium
was reached at station 3, and conditions stayed
close to equilibrium
up to the last sampling site. The waters were
undersaturated with
respect to smithsonite and cerussite at all
stations.
Conclusions.
The involvement of cyanobacteria in the formation
of calcium carbonate (11-13, 27) and other types of
carbonates (18, 23) is well documented. This precipitation
is helped by photosynthesis metabolism, which in the aqueous system
shifts the inorganic C species equilibrium to carbonates
(25). The biomineralization of zinc carbonates has not been
reported, except by Kalin (18), who did not specify the
mineralogical phase of the zinc carbonate.
Cell envelope characteristics are crucial for mineral nucleation
(
13). Since in natural conditions microbial organisms have
electronegative surfaces (
9), they easily bind metals to
their
envelopes. In our study several observations support the
epicellular
biomineralization of hydrozincite on cyanobacterial sheaths
induced
by alkalization in the microenvironment due to CO
2
fixation from
dissolved HCO
3 and release of
OH
during photosynthesis. In fact, an increase (rapid) in
pH associated
with a decrease in HCO
3 where
carbonate precipitation occurs is observed. The precipitation
of
minerals on cell envelopes generally results in a very fine
precipitate
(
24). In this study, for
Scytonema sp. strain
ING-1
we can propose a mechanism similar to the one proposed for a
cyanobacterial
Synechococcus sp. (
10,
26) and
include the following biological
and chemical reactions:
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(1)
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(2)
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(3)
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In spite of the acidity produced by reaction 3, the pH actually
increases, being abundantly counterbalanced by the high photosynthetic
activity.
Acidic polysaccharide (Alcian blue positive) binds Zn
2+ and
other metals. This metal binding, concomitant with
CO
32 formation during photosynthesis, causes
a local oversaturation
with respect to the carbonate phase, thus
favoring hydrozincite
precipitation.
The formation of hydrozincite instead of the anhydrous-phase
smithsonite reflects the chemical conditions imposed by biological
activity. Both these secondary minerals are common in a mine
environment.
Hydrozincite is stable for log pCO
2 >
5.2, whereas smithsonite
is stable at a higher level of
pCO
2 (log pCO
2 >
1.5) (
29).
The natural remediation of heavy metals produced by this photosynthetic
population can probably be exported to other similar
environments to
attenuate metal
pollution.
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ACKNOWLEDGMENTS |
This research work was supported by MURST and CNR grants.
We thank L. Fanfani and P. Lattanzi for their criticism of the paper.
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FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Environmental Sciences, University Cà Foscari, La Celestia, Via
Castello 2737/b, I-30122 Venice, Italy. Phone: 39-041-2578432. Fax:
39-041-5281494. E-mail: baldi{at}unive.it or
baldi{at}unisi.it.
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Applied and Environmental Microbiology, November 2000, p. 5092-5098, Vol. 66, No. 11
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