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Appl Environ Microbiol, April 1998, p. 1319-1322, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Biodegradation of Metal-EDTA Complexes by
an Enriched Microbial Population
Russell A. P.
Thomas,1
Kirsten
Lawlor,2
Mark
Bailey,2 and
Lynne E.
Macaskie1,*
School of Biological Sciences, The University
of Birmingham, Birmingham B15 2TT,1 and
NERC Institute of Virology and Environmental Microbiology,
Oxford OX1 3SR,2 United Kingdom
Received 26 August 1997/Accepted 9 January 1998
 |
ABSTRACT |
A mixed culture utilizing EDTA as the sole carbon source was
isolated from a mixed inoculum of water from the River Mersey (United
Kingdom) and sludge from an industrial effluent treatment plant.
Fourteen component organisms were isolated from the culture, including
representatives of the genera Methylobacterium,
Variovorax, Enterobacter,
Aureobacterium, and Bacillus. The mixed culture biodegraded metal-EDTA complexes slowly; the biodegradability was in
the order Fe>Cu>Co>Ni>Cd. By incorporation of inorganic phosphate
into the medium as a precipitant ligand, heavy metals were removed in
parallel to EDTA degradation. The mixed culture also utilized a number
of possible EDTA degradation intermediates as carbon sources.
| |
INTRODUCTION |
EDTA, an aminopolycarboxylic acid
chelating agent, has many applications. For example, it is used in
washing powders as a substitute for polyphosphates, which have been
implicated in the eutrophication of aquatic environments (11,
12). The large quantities of EDTA used commercially, and its
low-level biodegradability (11, 12), allow it to remain at
high levels in wastewater (11, 12, 14), from which it is not
removed by conventional wastewater treatment, and also to persist in
drinking water (10, 14, 36, 37). EDTA is also used for the
decontamination of nuclear power plant equipment (3, 20, 23,
31) because it forms very strong complexes with heavy metals and
makes them soluble and easy to remove from contaminated surfaces and
soils (22, 24). Conversely, if EDTA is not removed from
wastes before dumping, it can mobilize metals, e.g., the radionuclides
in nuclear waste dumps (8, 31). At the Oak Ridge National
Laboratory (Oak Ridge, Tenn.) and Maxey Flats (Kentucky) sites,
radionuclides were detected up to 100 m from the burial site
(8, 25), with EDTA-enhanced nuclide migration implicated in
nuclide mobilization (24, 25). Radionuclides normally bind
to humic and fulvic acids in soil and become immobilized due to the
tight complexes formed (7). EDTA prevents this, having a
higher affinity for the radionuclides than the humate and fulvate
ligands (7). It is now illegal to dump complexing agents
together with radionuclides in low-level burial sites in the United
States (29); such mixed wastes require pretreatment
(13). UV photodegradation of the ferric chelate of EDTA was
observed in early studies (16, 17), but extensive photolytic
mineralization does not occur (19) and this method of
degradation might not be applicable to large volumes. Alternatively,
biodegradation can release free metal, allowing this to be concentrated
via chemical or biotechnological methods (13).
EDTA is degraded slowly in the environment (6, 35). Its
biodegradation was proposed to comprise two pathways (4). The first involves the stepwise removal of acetate groups to leave ethylenediamine and the second involves the stepwise removal of an
iminodiacetate (IDA) group, a glycine, and then an ammonium group with
nitrilotriacetate (NTA)-aldehyde (4).
An Agrobacterium radiobacter strain able to biodegrade the
ammonium ferric complex of EDTA was isolated (18) but was
unable to degrade nickel, cobalt, or copper EDTA complexes or ferric complexes of NTA, IDA, or
ethylenediamine-N,N'-diacetic acid (EDDA) (18, 30), despite their role as possible intermediates of EDTA degradation (4). EDTA biodegradation by a mixed culture and one of its component organisms isolated from sewage sludge was
studied by Nortemann et al. (27, 28), using ferric ammonium EDTA as the sole nitrogen source initially and as the sole carbon source in later studies. A preliminary report suggested the possible growth of a mixed culture at the expense of heavy metal-EDTA complexes, but the loss of EDTA from the medium was not confirmed (21).
To date, the biodegradation of heavy metal complexes of EDTA, other
than Fe, has not been demonstrated. The present study investigated the
biodegradation of these complexes by an enriched microbial population
and suggested the possible use of such populations in the remediation
of wastes containing these complexes.
| |
MATERIALS AND METHODS |
Culture isolation and culture conditions.
Samples were taken
from the River Mersey (United Kingdom) and from liquid effluent sludge
from an industrial effluent treatment plant. The river water (8 ml) and
sludge (2 ml) were inoculated into 90 ml of filter-sterilized minimal
medium (MM) with the following components (grams per liter):
CaCl2, 0.025; MgSO4 · 7H2O,
0.2; NaCl, 0.1; (NH4)2SO4, 0.5;
disodium EDTA, 0.015; ZnSO4 · 7H2O, 0.0066; MnCl2 · 4H2O, 0.00171;
FeSO4 · 7H2O, 0.0015;
CoCl2 · 6H2O, 0.000483;
CuSO4 · 5H2O, 0.000471;
NaMoO4 · 2H2O, 0.000453;
3-(N-morpholino)propanesulfonic acid, 5.225; and
KH2PO4, 0.272. The pH was adjusted to 7 with 1 M NaOH, and 7.34 g of FeNaEDTA (Sigma, Poole, United Kingdom) liter
1 (20 mM) was added as the sole utilizable carbon
source. The cultures were shaken (160 rpm, 30°C) for a month and then
subcultured (10-ml inocula) into 90 ml of fresh MM with 1.83 g of
FeNaEDTA liter1 (5 mM); this was repeated monthly for 2 years.
To test for growth on heavy metal-EDTA complexes, samples (10 ml) taken
from cultures pregrown with FeNaEDTA as the sole carbon source (30 days) were harvested by centrifugation (7,000 rpm, MSE high-speed 18 centrifuge, room temperature), washed twice in sterile isotonic saline
(8.5 g of NaCl liter1), and inoculated into 100 ml of MM
containing 5 mM KH2PO4 and supplemented (to 5 mM) with CuNa2EDTA (1.88 g liter1),
CdNa2EDTA (2.22 g liter1),
CoNa2EDTA (1.96 g liter1),
NiNa2EDTA (1.96 g liter1) (all from TCI,
Tokyo, Japan), FeNaEDTA (1.83 g liter1), or disodium EDTA
(1.86 g liter1; Sigma) as the sole carbon source. The
EDTA compounds were of the highest grade commercially available (the
NaEDTA was 99% pure; the others were at least 96% pure, with sulfate
as the major contaminant, according to the manufacturer's
specifications), and the purity of each was confirmed by
high-performance liquid chromatography (see below). Controls were
samples without biomass (for each EDTA complex) and
non-EDTA-supplemented cultures. Cultures were incubated as described
above, with samples (1 ml) taken periodically and stored at 
20°C
for later analyses.
Growth on various substrates.
Samples (10 ml) were taken
from cultures previously grown on FeEDTA (30 days), harvested, and
washed as before and inoculated into MM (100 ml) supplemented with (5 mM) EDDA, ethylenediamine, trisodium acetate, trisodium NTA, IDA, or
glycine, with growth tested as described above. Controls were
uninoculated media and non-substrate-supplemented cultures.
Isolation and identification of microorganisms.
A
103 serial dilution was made in sterile isotonic saline
from a month-old FeNaEDTA culture. A sample (10 µl) was spread plated onto MM plates containing 2 mM sodium acetate (BDH, Poole, United Kingdom) or 2 mM FeNaEDTA (30°C). Isolates were Gram stained by the
Huckner method (9) with the Sigma Gram stain kit.
Gram-negative rods were identified with the API 20E and 20NE kits
(Biomerieux, Marcy l'Etoile, France). Isolates were also
identified by the Microbial Identification System (MIDI-MIS, Newark,
Del.) as described previously (33). Fatty acid methyl ester
(FAME) profiles were matched with those in the MIDI-MIS Trypticase soy
broth agar aerobic database (version 3.2).
Biomass immobilization.
Cultures (50 ml) previously grown on
1.83 g of FeEDTA liter1 (30 days), harvested, and
washed as described above were inoculated into 2.5 liters of MM
containing 1.5 g of ethylenediamine liter1 and
3.4 g of trisodium acetate liter1 (both 25 mM) in an
airlift reactor constructed in the laboratory. Shale particles (60 g;
average particle size, 3 by 3 mm; Thames Water) were washed with
distilled water, sterilized (10% Chloros, 2 days), and washed several
times with sterile distilled water. The particles were divided into two
30-g portions and placed into the lumens of two identical glass columns
(each with a 90-ml capacity; 14.5 by 2.8 cm). The culture was cycled
(Watson Marlow flow inducer) though both columns simultaneously at a
flow rate of 1 ml min1 for 5 days. Random particles from
each column were removed for protein analysis. After preparation, the
columns were stored at 4°C until analyzed. For EDTA degradation
experiments, 150 ml of MM containing 1 mM FeEDTA (0.367 g
liter1) and 1 mM KH2PO4 was
pumped through each column at a flow rate of 0.25 ml min1
(single pass). Column outflow samples, withdrawn hourly, were stored at
20°C for later analysis, together with samples of the input
solution.
Analyses of biomass, residual EDTA, and metals.
Frozen
samples (1 ml) were thawed, the biomass was removed by centrifugation
(13,000 rpm in a Heraeus Sepatech Biofuge A at room temperature), and
supernatant samples were removed, filtered (0.45-µm pore size), and
diluted into fresh vacuum-filtered and degassed MM. EDTA analysis was
based on a previously described method (5) using
high-performance liquid chromatography with a 410 series UV detector
(the pumps, injector, and detector were all from Waters, Milford,
Mass.) and a Nucleosil 5-µm C18 column, 250 by 4.6 mm
(Phenomenex, Macclesfield, Cheshire, United Kingdom). The mobile phase
was 0.03 M sodium acetate/acetic acid buffer (pH 4)/20 mM tetrabutyl
ammonium hydroxide and 100 ml of methanol liter1,
filtered and degassed as described above. EDTA was detected at
wavelengths of 254 and 300 nm. Metals were assayed by atomic-absorption spectroscopy (Pye Unicam, Cambridge, United Kingdom), with an acetylene
(9-lb/in2) and air (30-lb/in2) flame. Cd, Cu,
Zn, Ni, and Co were detected at wavelengths of 228.8, 324.8, 213.9, 232.0, and 240.7 nm, respectively, against a blank of fresh MM.
For protein analysis, cell pellets were resuspended in 50 mM NaEDTA (1 ml), mixed for 10 min to remove bound metals, recentrifuged, and
resuspended in MilliQ water. Protein was measured by the copper sulfate-bicinchoninic acid method (reagents were all from Sigma). For
protein analysis of biomass-loaded shale, specimen shale particles were
placed in a 1.5-ml tube containing MM with 50 mM NaEDTA (500 µl) and
mixed vigorously for 10 min. This was repeated until no protein was
detected in the wash solution. The dislodged cells were harvested and
the protein concentration was estimated as described above.
Treatment of results.
All experiments were done in
triplicate on three separate occasions, and the data were pooled and
calculated as means ± standard deviations (SD) for three
replicates. In general the SD was within ±10% of the mean and is only
shown where this value was exceeded. Means ± SD for individual
values are given below as appropriate.
| |
RESULTS AND DISCUSSION |
Microorganism isolation and identification.
Fourteen
morphologically distinct colony types were isolated from the mixed
culture after 18 subcultures. Four were gram-negative bacteria and were
identified by the API 20E and 20NE systems as strains of
Enterobacter and Pseudomonas, respectively.
These, and the 10 gram-positive bacteria, were identified further on the basis of cellular FAME profiles. Of the culturable organisms within
the culture, gram-positive bacteria predominated, comprising species of
Methylobacterium (35%), Variovorax (17%),
Bacillus (10%), and Aureobacterium (10%) (Table
1). Variovorax
(Alcaligenes) paradoxus was shown previously to
tolerate EDTA (15) or, possibly more correctly, the Fe
starvation that could be imposed by extensive tight chelation of Fe.
This organism can degrade a variety of xenobiotics and also
hydroxybutyrate, an ability it shares with Aureobacterium
saperdae (2, 26, 32, 37, 38), which was also found in
the present culture (Table 1). The microorganisms identified in Table 1
represent only those that were culturable on plates. It is likely that
a number of other microorganisms contributing to EDTA biodegradation
but probably unculturable in isolation were also present. The total
number of microorganisms found per milliliter was 10- to 100-fold less
than expected on the basis of viable counts and compared with the
determined amount of protein per milliliter. Therefore, it can be
assumed that >90% of the microorganisms are not culturable in
isolation by the method used here. This is supported by a published
report which suggests that <1% of all microorganisms are culturable
(1). Further studies using molecular probe methods are
warranted.
Biodegradation of metal-EDTA complexes and likely metabolic
intermediates.
Incubation of the culture with the metal-EDTA
complexes resulted in growth, biodegradation of each complex, and
removal of metal over 28 days (Fig. 1).
Equimolar inorganic phosphate was incorporated to scavenge released
metal by precipitation, avoiding metal toxicity and illustrating the
potential for remediation of metal-EDTA wastes. Growth on Fe- and
NaEDTA gave doubling times of 66 and 288 h, respectively, with
corresponding levels of EDTA degradation of 3.01 ± 0.19 mM (60%)
and 1.57 ± 0.26 mM (31%) after 28 days (Fig. 1B and A).
Biodegradation of the FeEDTA complex also resulted in the removal of
44% ± 0.03% of Fe from solution, probably as the hydroxide or
phosphate. The apparently poor growth with and low biodegradability of
NaEDTA in comparison to FeEDTA are probably due to the replacement of
the Na ligand with essential trace metals from the medium, effectively
starving the cells of these metals. Growth at the expense of the
recalcitrant heavy metal complexes, Cd-, Cu-, Co-, and NiEDTA, was
observed. The respective extents of EDTA biodegradation for these were
0.95 ± 0.13 mM (19.3%), 1.54 ± 0.23 mM (30%), 1.3 ± 0.23 mM (25%), and 1.16 ± 0.24 mM (23.4%) (Fig. 1). The
corresponding extents of metal removal were 19% ± 0.02% (Cd), 16% ± 0.01% (Cu), 14% ± 0.01% (Co), and 31% ± 0.00% (Ni) (Fig. 1C
to F). Metal removal was stoichiometric or greater in the cases of Cd
and Ni but only approximately 50% of that released from EDTA with Cu
and Co. Insufficient metal was precipitated to permit X-ray powder
diffraction analysis of the recovered precipitate.
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FIG. 1.
Growth ( ) of the mixed culture on MM supplemented
with 5 mM inorganic phosphate and NaEDTA (A), Fe(III)EDTA (B), CdEDTA
(C), CuEDTA (D), CoEDTA (E), and NiEDTA (F) complexes; EDTA consumption
( ); and metal removal ( ). All SD are within 10% except where
indicated.  , control (non-substrate-supplemented culture).
|
|
From the above data the biodegradability of the heavy metal-EDTA
complexes was ranked as follows: Fe>Cu>Co>Ni>Cd.
From previous
studies it could be predicted that FeEDTA would be the
most readily
biodegraded (
4,
18). Previous work
(
18) also demonstrated
biodegradation of Fe(III)EDTA by an
Agrobacterium sp. at a rate
of approximately 275 µmol
h
1 µg of protein
1, which was nearly
20-fold higher than that obtained with the
present mixed culture (15 µmol h
1 µg of protein
1). With a mixed
culture, not all of the component organisms will
degrade EDTA per se
and may instead scavenge low-molecular-weight
products, contributing to
the total biomass protein and lowering
the specific activity against
the parent compound. The above-mentioned
Agrobacterium sp.
was of limited potential, being unable to biodegrade
EDTA at
concentrations below 5 mM or to utilize other metal-EDTA
complexes.
However, in addition to being versatile, the present
culture gave a
residual EDTA concentration of 2 to 2.5 mM (FeEDTA),
with continuing
activity at termination of the experiment at 30
days. Degradation of
the other heavy metal-EDTA complexes appeared
to cease at 3 to 4 mM
EDTA (Fig.
1). This could be due to a specific
threshold (e.g., as for
the
Agrobacterium sp.) below which the
complexes of metals
other than Fe are unable to be transported
into the cells; the biphasic
nature of the removal of FeEDTA (Fig.
1B) could suggest the presence of
two uptake processes, one nonspecific,
low-affinity system and one
Fe-specific, higher-affinity, system.
The apparent
Km values for the various complexes have never
been
studied and would repay further investigation, as they may
ultimately
limit the usefulness of an industrial process. However,
such
Km data would be difficult to interpret by
using a mixed culture
because individually, the isolates grew poorly at
the expense
of EDTA.
Other metal complexes (Mg-, Mn-, and CaEDTA) have been biodegraded
completely by both a mixed microbial population and a single
isolate.
However, these metals are of relatively low toxicity
and are essential
micronutrients (
27). A study of CoEDTA biodegradation
using
an Fe(III)EDTA-biodegrading
Agrobacterium sp. was
successful,
but only because of displacement of the Co by Fe(III)
(
30).
Biodegradation of the Co, Cu, Ni, and Cd complexes of
EDTA has
not been reported previously. Other workers have suggested the
role of a specific Fe binding siderophore in the biodegradation
of
Fe(III)EDTA (
18), and indeed, EDTA degradation did not cease
as with the other metals but continued at a lower rate after 5
days
(Fig.
1). It has been suggested that microorganisms isolated
from an
EDTA-rich environment have developed a specific Fe transport
mechanism
which is dependent on the binding of EDTA (
18).
Previous studies have paid little attention to the likely breakdown
intermediates of EDTA as potential "bottlenecks" which
may limit
the potential for its complete mineralization. The culture
in the
present study grew on substrates (EDDA, ethylenediamine,
trisodium
acetate, NTA, IDA, and glycine) previously proposed
as degradation
intermediates (
4). Growth was more rapid than
on FeEDTA for
all substrates except EDDA, which, after a lag,
gave a growth rate
equal to that on EDTA (Fig.
2). Buildup
of
these intermediates was therefore unlikely to inhibit complete
EDTA
mineralization.
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FIG. 2.
Growth of the culture on acetate ( ), ethylenediamine
( ), IDA ( ), glycine ( ), NTA (), EDDA ( ), and FeEDTA
(). , control (non-substrate-supplemented culture). All SD are
within 10%.
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|
EDTA biodegradation by immobilized cells.
The potential for
developing a continuous-flow method of treating wastes bearing
metal-EDTA complexes was assessed by using biomass samples of the mixed
culture immobilized as a biofilm on shale particles. At a residence
time of only 2.5 h (the fluid volume was 37 ml; the flow rate was
0.25 ml min1), the immobilized cells degraded 20% ± 0.02% of the supplied FeEDTA and 13% ± 0.03% of the Fe was removed
(150 ml of 1 mM FeEDTA over 10 h). In contrast, the time required
for 20% removal of FeEDTA in the batch system was 4.5 days (Fig. 1).
The input concentration of EDTA here was 1 mM, below the concentration
which could be degraded by Agrobacterium in previous studies
(see above).
The generation of a natural biofilm on shale particles may not be the
most suitable method for the continuous treatment of
metal-EDTA wastes.
As an alternative, immobilization onto microcarriers
as described
previously (
34) could be a superior method, and
this should
be investigated in future studies.
In conclusion, we demonstrate for the first time the feasibility of
biodegradation of several heavy metal-EDTA chelates and
also identify
some unusual microorganisms implicated in this process.
Significant
removal of EDTA in a continuous-flow system by immobilized
biofilm
justifies development of process engineering aspects of
the system and
further elucidation of the microbiological composition
and biochemical
processes within the culture.
| |
ACKNOWLEDGMENTS |
We thank Thames Water for the gift of the shale particles and G. Basnak for help with the analysis of metals by atomic-absorption spectroscopy.
The support of the BBSRC (studentship no. 93302170 to R.A.P.T.) is
acknowledged, with thanks. K.L. was supported by a grant from the
European Union (EU Bio2 CT-943001).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: School of
Biological Sciences, The University of Birmingham, Edgbaston,
Birmingham, B15 2TT, United Kingdom. Phone: (44) 121-414-5889. Fax:
(44) 121-414-6557. E-mail: L.E.Macaskie{at}bham.ac.uk.
| |
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Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
