Open Journal of Metal, 2013, 3, 51-61 Published Online July 2013 (
Apatite and Chitin Amendments Promote Microbial
Activity and Augment Metal Removal in
Marine Sediments
Jinjun Kan1*, Anna Obraztsova2, Yanbing Wang3, Jim Leather2, Kirk G. Scheckel4,
Kenneth H. Nealson3, Y. Meriah Arias-Thode2*
1Stroud Water Research Center, Avondale, USA
2SPAWAR Systems Center-Pacific, San Diego, USA
3Department of Earth Sciences, University of Southern California, Los Angeles, USA
4National Risk Management Research Laboratory, Environmental Protection Agency, Cincinnati, USA
Email: *,
Received May 4, 2013; revised June 12, 2013; accepted June 19, 2013
Copyright © 2013 Jinjun Kan et al. This is an open access article distributed under the Creative Commons Attribution License,
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
In situ amendments are a promising approach to enhance removal of metal contaminants from diverse environments
including soil, groundwater and sediments. Apatite and chitin were selected and tested for copper, chromium, and zinc
metal removal in marine sediment samples. Microbiological, molecular biological and chemical analyses were applied
to investigate the role of these amendments in metal immobilization processes. Both apatite and chitin promoted micro-
bial growth. These amendments induced corresponding bacterial groups including sulfide producers, iron reducers, and
phosphate solubilizers; all that facilitated heavy metal immobilization and removal from marine sediments. Molecular
biological approaches showed chitin greatly induced microbial population shifts in sediments and overlying water: chi-
tin only, or chitin with apatite induced growth of bacterial groups such as Acidobacteria, Betaproteobacteria, Epsilon-
proteobacteria, Firmicutes, Planctomycetes, Rhodospirillaceae, Spirochaetes, and Verrucomicrobia; whereas these
bacteria were not present in the control. Community structures were also altered under treatments with increase of rela-
tive abundance of Deltaproteobacteria and decrease of Actinobacteria, Alphaproteobacteria, and Nitrospirae. Many
of these groups of bacteria have been shown to be involved in metal reduction and immobilization. Chemical analysis
of pore- and overlying water also demonstrated metal immobilization primarily under chitin treatments. X-Ray absorp-
tion spectroscopy (XAS) spectra showed more sorbed Zn occurred over time in both apatite and chitin treatments (from
9% - 27%). The amendments improved zinc immobilization in marine sediments that led to significant changes in
the mineralogy: easily mobile Zn hydroxide phase was converted to an immobile Zn phosphate (hopeite). In-situ
amendment of apatite and chitin offers a great bioremediation potential for marine sediments contaminated with heavy
Keywords: Apatite; Chitin; Amendments; Marine Sediment; DGGE; Microbial Community; Copper; Zinc; Chromium
1. Introduction
In situ amendments have been shown as an effective ap-
proach to enhance removal of metal or organic contami-
nants under a variety of environments [1-6]. Currently,
both inorganic and organic materials are widely used as
promising sediment amendments; including geotextile
mats, apatite, organoclay, and chitin. Inorganic amend-
ments help to immobilize several toxic metals such as
lead, zinc, and cadmium by forming stable compounds
that render the metals no longer bioavailable [2,3,7]. Or-
ganic amendments, in contrast, may induce indigenous
microbes capable of bio-reduction and/or of direct or
indirect immobilization of toxic metals [1,4,8,9].
Apatite, an inorganic phosphate mineral, has been ap-
plied to facilitate the immobilization of several toxic
metals including Cu, Zn and Pb [10-12]. For example,
apatite amendments have a long remedial history at met-
als contaminated soils sites [10,13,14]. Apatite’s use in
sediment remediation is more recent [15]. The DoD
(Navy) is currently evaluating apatite amendments at a
*Corresponding author.
opyright © 2013 SciRes. OJMetal
metal recycling facility to remediate high Zn levels in
nearby sediments [16]. Phosphates released from apatite
will sequester metals to form more stable metalphosphate
complexes [3]. This process may be linked to activities
of phosphate-solubilizing bacteria. Although marine en-
vironments are not phosphate-limited, recently Uzair and
Ahmed [17] have successfully isolated both attached and
free-living marine bacteria that were able to solubilize phos-
phate compounds when given suitable carbon sources.
Meanwhile chitin, a homopolymer of N-acetylgluco-
samine (NAG), is an abundant structural polysaccharide
produced by many marine organisms. Several experi-
ments have demonstrated that bacterial communities shift
in response to amended chitin [18] and indigenous bacte-
ria were capable of cleaving this polymer via the synthe-
sis of specific enzymes that were induced by chitin [19].
Further, breakdown products (e.g. acetate and fructose)
from chitin degradation may serve as ideal carbon sources
for anaerobic and facultative microorganisms capable of
metal reduction [20]. It is thus reasonable to expect that
we hypothesize that amendment of apatite and/or chitin
will stimulate bacterial growth, and subsequently en-
hance metal transition, immobilization and sequestration
in marine environments.
Metabolically diverse indigenous microorganisms in-
teract with metals in a variety of ways that lead to de-
creased metal solubility and mobility [21-22]. Among
these microbes, iron-reducing and sulfide-producing bac-
teria (FeRB and SPB, respectively) are two biogeo-
chemically important groups that have been shown to
contain suitable physiology for metal precipitation and
immobilization due to their metabolic end-products such
as Fe(II) and HS [23-29].
In addition to the effectiveness of bioremediation, one
other critical perspective of inorganic or organic materi-
als used as in situ amendment remediation alternatives, is
to limit/avoid the impacts on living organisms. Therefore,
during our primary study, we tested how the inorganic
and organic amendments impacted microbial communi-
ties [30] and impacted the macro-invertebrate communi-
ties [31]. Dose-response experiments of candidate mate-
rials suggested that appropriate concentrations of apatite
and chitin (5% and 0.5%, respectively) amendments had
little or no negative impacts on water quality and ambient
living organisms [31].
In this study, apatite and chitin were applied to micro-
cosms containing pristine, coarse, sandy sediments from
Yaquina Bay (YB), Oregon, and contaminated, fine-
grained sediments from Mare Island (MI) Naval Ship-
yard, California, with and without spiked heavy metals.
Responses of the microbial communities (biomass and
population structures) were monitored in the treated sedi-
ments. Chemical analysis, including ICP-MS and X-ray
absorption spectrometry was used to determine the ef-
fects of the amendments on the sediment composition
and zinc speciation.
2. Experimental
2.1. Sediment Description
Pristine and uncontaminated, sandy, coarse sediments
from Yaquina Bay (YB), OR, and historically heavy
metal contaminated, fine-grained sediments from Mare
Island (MI) Naval Shipyard, CA, were evaluated [29].
Major properties of both sediments including percentage
of silt/clay, TOC (total organic carbon), and metals were
listed in a previous study [31]. Because the MI sediments
were aged, additional spikes of Cu, Zn, and Cr were in-
corporated with the goal of metal concentrations in the
overlying water of Cu = 0.25 ppm, and Zn and Cr = 1.0
ppm (Table 1). These concentrations were used as a sur-
rogate for current metal contaminated sites. The metal
additions were added with the assumption of no binding
as TOC of coarse sand is less than 0.1%. The sediment
types and treatments applied in this study are listed in
Table 1.
2.2. Amendment Experiments with Apatite
and Chitin
Mesocosm preparation and experimental setups followed
previously described methods [30-31]. Briefly, for each
treatment, 150 g of sediment was added to each of 5 rep-
licate beakers. A sixth replicate was used for monitoring
daily water. Approximately 750 mL of uncontaminated
and filtered (0.45 μm Millipor filter) natural seawater
(salinity at 30 psu) collected from near the mouth of San
Diego Bay, was added to each jar. A 3-day equilibration
period preceded the experiments. All beakers were gently
and continuously aerated.
Based on the previous toxicological dose-response
experiments and microbial response studies [30-31], apa-
tite (5%) and chitin (0.5%) were selected as candidate
amendment treatments. Concentrations of amendments
and treatment setups are described in Table 1. Negative
controls contained unspiked and unamended YB and MI
sediments (Table 1; CUU and FUU, respectively).
2.3. Microbiological Sample Collection and
Water and sediment core samples were collected for all
analyses at certain time points as required. Because many
metal-microbe interactions occur in the overlying water
and surface sediments, overlying water and pore-waters
were analyzed for bacterial populations. Detailed de-
scriptions of sample collections and treat ments are pre-
viously described [30]. Samples for molecular and mi-
crobiology analyses were collected at Day 0 and Day 28.
Copyright © 2013 SciRes. OJMetal
J. KAN ET AL. 53
Table 1. Sediment amendments applied in this study.
type Amendment Metal mixture*
Coarse (YB) Unamended Unspiked CUU
Coarse (YB) Unamended Spiked CUS
Coarse (YB) 5% apatite Spiked CAS
Coarse (YB) 0.5% chitin Spiked CCS
Coarse (YB) 5% apatite + 0.5% chitinSpiked CACS
Fine (MI) Unamended Unspiked FUU
Fine (MI) 5% apatite Unspiked FAU
Fine (MI) 0.5% chitin Unspiked FCU
Fine (MI) 5% apatite + 0.5% chitinUnspiked FACU
Fine (MI) Unamended Spiked FUS
Fine (MI) 5% apatite Spiked FAS
Fine (MI) 0.5% chitin Spiked FCS
Fine (MI) 5% apatite + 0.5% chitinSpiked FACS
YB: Yaquina Bay, OR; MI: Mare Island Naval Shipyard, CA; *Appropriate
salt forms of Cu, Zn, and Cr were incorporated into the overlying water
column to achieve free metal concentrations of Cu = 0.25 ppm, and Zn and
Cr = 1.0 ppm.
2.3.1. Microbiological Analysis
Microbial biomass was determined by epifluorescence
microscopy. DNA extraction, PCR-DGGE (denaturant
gradient gel electrophoresis) of 16 S ribosomal RNA
gene, band excision and sequencing, and phylogenetic
analyses followed described protocols in a previous re-
port [30].
2.3.2. Clone Library Analysis
Based on the fingerprinting results from PCR-DGGE,
and to further characterize the bacterial communities
from sediments, three clone libraries were constructed
from the anaerobic zone on the treatments FUS, FCS,
and FACS (Table 1). This analysis could not be com-
pleted from the coarse-grained sandy sediments because
of insufficient DNA concentrations. Briefly, near full
length of 16 S rRNA genes was amplified with primer 27
F-1492 R, and PCR products from triplicate samples
were pooled to minimize variations. Then the amplicons
were cloned using the TOPO TA Cloning Kit (Invitrogen)
and then sequenced by using the Big Dye Terminator
chemistry (Applied Biosystems). Chimeric sequences were
detected using the program of Bellerophon [32] and re-
moved from further analysis. All non-chimeric sequences
were blasted against the GenBank (http://www.ncbi.nlm. Sequences from both DGGE bands
and clone libraries were submitted to GenBank under the
accession number KF268681-KF268939.
2.3.3. Phosphate Solubilizing Bacteria Detection
Three dilutions (101, 102, and 103) of overlying water
and sediments (aerobic/anaerobic interface) were tested
to detect phosphate-solubilizing bacteria on apatite agar
plates. Plates were prepared as previously described [33]
with some modifications, i.e. 6 g CaCl2 and 4 g KH2PO4
were added to 500 mL filtered (0.22 µm Millipore™)
seawater. To a separate 500 mL of sterile seawater, 10 g
agar was added and all solutions were autoclaved sepa-
rately. 20 mL of 1 M sterile NaOH and 20 mL of 1 M
sterile glucose was added to the CaCl2/KH2PO4 seawater
solution and mixed well. After that, the CaCl2/KH2PO4
seawater solution wasmixed with autoclaved agar solu-
tion and poured into plates. Diluted samples from the
overlying water column and sediments were inoculated
and streaked for isolation on the agar plates; incubated
for 2 weeks at room temperature.
2.4. Chemical Analyses
Analyses for overlying and pore-water samples by ICP-
MS were taken at Day 0 and Day 10, due to the high mi-
crobial activity. Sediments were examined via X-ray
absorption spectroscopy (XAS) at Day 0 and Day 10. To
examine long-term effects of apatite, a few selected sam-
ples were analyzed at Day 28, and one sample from the
unspiked apatite amendment was extended to 120 days.
2.5. ICP-MS Analysis
Metal extraction from overlying and pore-water followed
EPA sample preparation protocol and metal analysis with
ICP-MS were performed as previously described [34-35].
2.6. X-Ray Absorption Spectroscopy (XAS)
XAS analyses followed the exact protocol from a previous
report [30]. Briefly, the samples were prepared as thin pel-
lets with a hand operated IR pellet press and the samples
were secured by Kapton tape. Five individual spectra for
each sample were averaged followed by subtraction of the
background through the pre-edge region using the Autobk
algorithm and normalized to an atomic absorption of one.
The data were converted from energy to photoelectron
momentum (k-space) and weighted by k3. Identification
of zinc phases in the sediment samples was accomplished
by principal component analysis (PCA) and linear com-
bination fitting (LCF) of the sediment XAS spectra rela-
tive to the known reference spectra. Reference materials
examined include hopeite (Zn3(PO4)2·4H2O), Zn-Al lay-
ered double hydroxide with nitrate and silicate interlayers,
Zn(OH)2, ZnO, sphalerite (ZnS), zinc sorbed to ferrihy-
drite, zinc sulfate, aqueous zinc nitrate, franklinite
(ZnFe2O4), willemite (Zn2SiO4), hemimorphite
(Zn4Si2O7(OH)2·H2O), and gahnite (ZnAl2O4). The accu-
Copyright © 2013 SciRes. OJMetal
racy of LCF results was estimated to be 10% - 15% [36].
3. Results and Discussion
3.1. Microbial Population Analysis
Total chitin production by crustaceans in marine envi-
ronments are estimated to reach up to 109 t per year [37],
and most of the chitin (90%) is digested within 150 h in
water column [38]. In marine sediments, chitin degrada-
tion is slower and the breakdown process is primarily
accomplished by indigenous bacteria with capability of
digesting chitin extracellularly [39]. Therefore, it is ex-
pected that high concentrations of chitin will favor
growth and enrichment of certain bacterial groups. In this
study, substantial increases of microbial cell numbers as-
sociated with chitin amendments were observed after a
28-day incubation (Figure 1). Epifluorescence microsco-
pic counts showed an increase in cell numbers in overly-
ing waters from all treatments containing chitin: CCS,
CACS, FCU and FCS. The numbers were from 0.38 ×
106 cells/mL (control) to 3.27 × 106, 1.85 × 106, 1.08 ×
106, 2.42 × 106 cells/mL, respectively. However, cell den-
sity in the fine-grained sediments in the apatite plus chi-
tin amendments, FACU and FACS, did not demonstrate
statistically significant growth compared to the control.
Chitin has been widely used as a bio-control material
in soils in order to reduce parasitic diseases. Previous
reports showed that chitin amendment was associated
with an increase of bacterial populations with chitinolytic
activities such as actinomycetes [40-41]. Major break-
down products from chitin are acetate and fructose [20],
both of which may serve as carbon sources for anaerobic
and facultative microorganisms. Recently, Kanzog and
colleagues [18] reported that both microbial numbers and
chitinolytic activity significantly increased in the sedi-
ment samples enriched with chitin in an experiment con-
ducted in deep-sea sediment, indicating that chitin en-
hanced microbial growth, enzymatic activity, and popu-
lation structures, as well. Therefore, chitin may play an
important role by providing a slow-release carbon source
to maintain microbial activity.
3.2. Microbial Community Structure
Several studies have already shown that microbial com-
munity structures respond to variations of organic amend-
ments such as chitin [18,40,41]. PCR-DGGE in combi-
nation with DNA sequencing aided in identifying the
predominant bacterial populations and to provide a com-
prehensive picture of microbial community structure
over time and under different amendment and metal con-
ditions. Our study provides a “snapshot” for shift of bac-
terial population structures under chitin amendments.
Analysis of DGGE band profiles has indicated that
Figure 1. Microscopic cell counts in overlying waters after
28 days incubation. X axis showed the treatments as listed
in Table 1. *Significant difference compared to the control
(unpaired t-test, p < 0.05). Data from treatments CAS, FAU,
and FAS were not available.
chitin amendments shifted the bacterial population struc-
tures significantly in overlying waters (Figure 2). Phaeo-
bacter (band 1) and Roseobacter (band 2) were the pre-
dominant groups in the controls (CUU and FUU), while
more distinct bacterial groups were present after chitin
and/or apatite treatments in both sediment types (CCS,
CACS, FCU, FACU, FCS and FACS, Figures 2 and 3).
Most of these enriched bands (No. 5, 6, 8, 9, 10, 12, 14,
15, etc.) belonged to subgroups of Alphaproteobacteria
(Figures 2 and 3). However, Betaproteobacteium (Band
No. 3 Figure 3), was identified in most samples (CUU,
CUS, CAS, FAU, FUS, and FCS). In fact, DGGE profiles
from apatite and/or spiked metals were remarkably simi-
lar to those from control sediments. Therefore, compared
to chitin, addition of apatite or metals had less impact on
the bacterial population structures (Figures 2 and 3).
For sediments, more distinct and diverse bacterial
groups were enriched in both aerobic/anaerobic interface
and anaerobic zone sediments (Figure S1). As an exam-
ple, in coarse-grained (YB) surface sediments, Bacter-
oidetes (bands 19, 20), Spirochaetes (band 21), and Ro-
seobacter from Alphaproteobacteria (band 22) were pre-
sent in both chitin, and chitin + apatite treatments (CCS
and CACS, Figures S1(a) and S2). In contrast, Del-
taproteobacteria (bands 27, 28) and Bacteroidetes (bands
25, 26) dominated in anaerobic zone sediments from the
same treatments (CCS and CACS, Figures S1(b) & S2).
Due to the detection limit and sensitivity of DGGE ap-
proach, treatments from fine sediments (MI) didn’t gen-
erate distinguishable banding patterns and therefore, we
further characterized bacterial community structures by
clone library.
As we expected, remarkable population shifts were
observed from clone library analyses (Table 2 and Fig-
ure 4). Actinobacteria, Alphaproteobacteria, Gam-
maproteobacteria, Deltaproteobacteria, and Bacteroide-
tes were predominant groups in unamended fine-grained
sediments with spiked metals (FUS). After 28 days in-
Copyright © 2013 SciRes. OJMetal
J. KAN ET AL. 55
Table 2. Bacterial population structures characterized by
clone library. Relative abundance was based on the occur-
rence of representative sequences. - = not detected.
Relative abundance (%)
Acidobacteria - - 1.9 Induced
Actinobacteria 20.8 8.6 3.8 Decreased
Alphaproteobacteria 22.9 14.3 13.2 Decreased
Betaproteobacteria - 2.9 1.9 Induced
Gammaproteobacteria 22.9 10 20.8 Decreased
Amphritea 2.1 - 1.9
Marinobacterium 2.1 - -
Oceanospirillum 14.6 1.4 -
Paramoritella - - 3.8
Pseudomonas 2.1 - -
Shewanella - - 1.9
Uncultured 2.1 8.6 13.2
Deltaproteobacteria 14.6 30 28.3 Decreased
Desulfotalea - 1.4 3.8
Desulfovibrio - 5.7 3.8
Desulfuromonas - 5.7 -
Desulfobacula - 1.4 1.9
Geobacter - 1.4 -
Pelobacter 2.1 - -
Uncultured 12.5 14.3 18.9
Epsilonproteobacteria - 4.3 9.4 Induced
Bacteroidetes 10.4 10 11.3
Chloroflexi 2.1 1.4 1.9
Firmicutes - 2.9 5.7 Induced
Nitrospirae 6.3 4.3 - Decreased
Planctomycete - 5.7 - Induced
Rhodospirillaceae - - 1.9 Induced
Spirochaeta - 4.3 -
Verrucomicrobia - 1.4 - Induced
cubation with chitin and chitin + apatite, distinct bacterial
groups were induced, including Acidobacteria, Betapro-
teobacteria, Epsilonproteobacteria, Firmicutes, Planc-
tomycetes, Rhodospirillaceae, Spirochaetes, and Verru-
comicrobia, which were not detected in the unamended
treatment FUS (Table 2). To date, it is not easy to link
the functionality of these bacterial groups with their ge-
netic identification. However, high occurrence of these
groups of microorganisms under heavy metal concentra-
tions, e.g. Firmicutes indicated the potential direct or
Figure 2. DGGE fingerprints of bacterial community from
overlying waters (W). Labels referred to the treatments.
Sample FAS failed to obtain PCR products. Bands 1-15
were selected and excised for sequencing.
haeum equitans A
DGGE band 3
Uncultured betaproteobacterium GQ34109
DGGE band 8
Uncultured bacterium FN421572
Uncultured bacterium FJ202706
Uncultured bacterium EU183981
DGGE band 7
Uncultured Terasakiella sp. FJ753149
DGGE band 6
Donghicola eburneus DQ667965
Uncultured bacterium EF574092
Donghicola sp. EF587950
DGGE band 12
Uncultured marine bacterium AF177550
DGGE band 1
Phaeobacter sp. AB498882
DGGE band 13
Uncultured bacterium GU118445
Uncultured bacterium GU119459
DGGE band 15
Maritimibacter sp. EU052764
Uncultured alphaproteobacterium AY145603
Uncultured bacterium AJ319859
Psdeudoruegeria sp. FJ374173
Uncultured alphaproteobacterium GQ204861
DGGE band 9
DGGE band 14
Uncultured bacterium GU066436
DGGE band 10
Rhodobacteraceae bacterium AM990789
Rhodobacteraceae bacterium FJ460070
DGGE band 5
Uncultured alphaproteobacterium AM850968
DGGE band 2
DGGE band 4
DGGE band 11
Marine bacterium AF359535
Roseobacter sp. AY576690
Figure 3. Phylogenetic analysis (DNA distance-NJ) of bac-
terial DGGE band sequences obtained from overlying wa-
ters. Bootstrap values were based on analyses of 1000 re-
sampling of dataset. Nanoarchaeumequitans was used as an
outgroup. Scale bar represent 0.05 substitutions per site.
indirect involvement with metal immobilization or bio-
remediation [42-43]. Further, over time Deltaproteobac-
teria population increased in abundance, while that of
Actinobacteria, Alphaproteobacteria, Nitrospirae de-
creased (Table 2 and Figure 4). In fact, Deltaproteobac-
teria became the most dominant groups of bacteria after
Copyright © 2013 SciRes. OJMetal
Figure 4. Clone library analyses of bacterial population
structures from treatments FUS, FCS and FACS.
28 days incubation in both FCS (30%) and FAC (28.3%)
(Table 2). Predominant sulfate-reducing bacteria (i.e.
Desulfovibrio, Desulfobacula, Desulfotalea) and sulfur-
reducing bacteria (i.e. Desulfuromonas) sequences were
detected in Deltaproteobacteria class. These results agreed
with previous observations on increasing sulfate-reduc-
ing bacterial communities and activities in response to
carbon source amendments [44]. Primarily due to their
metabolic end-products such as sulfide, sulfate/sulfur re-
ducing bacteria were able to facilitate metal sequestration
and immobilization [24,45]. In addition, these bacteria
have been noted for their capability of utilizing metals as
electron acceptors via dissimilatory metal reduction [28-
29]. A good example is Geobacter, which was also re-
trieved from our clone library analyses from FCS (Table
2). In line with Geobacter, Shewanella of the Gam-
maproteobacteria class, another recognized metal re-
ducer [46], was also present in the treatment with chitin +
apatite (FACS). Both Geobacter and Shewanella have
been proven to play critical roles in metal reduction, pre-
cipitation and immobilization [1,23,24,27,45]. Thus, sul-
fate/sulfur-reducing bacteria and metal-reducing bacteria
responded to chitin and apatite amendments, and were
likely responsible for the metal immobilization and se-
Although no significant bacterial cell counts and popu-
lation shifts were observed in the treatment with apatite
alone [30 and this study], apatite held great promise of
inducing phosphate-solubilizing bacteria (PSB), which
might facilitate the formation of phosphorites, natural
apatite mineral deposits in environments [47-49]. Transi-
tion metals in natural environments were expected to be
sequestered with phosphates and form more stable metal-
phosphate complexes and therefore decrease the bioavail
ability of these metals [3]. In this study, we tested PSB
on agar plates, and positive PSB colonies were present in
both overlying water and sediments (Figure S3). The
clearing zones indicated solubilization of calcium phos-
phate from the media (Figure S3). To date, a variety of
bacterial groups (e.g. Bacillus, Rhizobium, Pseudomonas,
Serratia, Shewanella, Escherichia, Vibrio, and Proteus
etc.) have been proven capable of solubilizing phosphate
compounds [17,50-51]. Because phosphate is not a lim-
iting factor in open oceans, only few studies have inves-
tigated PSB from marine environments [33,51]. However,
the presence of common groups of bacteria including
Pseudomonas, Shewanella, and Vibrio from our clone
libraries suggested that they might respond to apatite
amendment and solubilize phosphate in marine sedi-
3.3. ICP-MS Analysis of Spiked Metals
In the presence of apatite only, less than 50% of Cu and
Cr; and 0% of Zn were removed from the overlying wa-
ter in the coarse-grained sediments (CAS; Table 3). A
similar effect was observed in the fine-grained sediment
as Cu and Cr were not removed at all; and in the Zn
sample, ~25% of Zn was removed (FAS; Table 3). In the
presence of chitin, between 60% - 100% of metal re-
moval occurred in the overlying and pore-water (CCS
and FCS; Table 3). The combined effect of apatite and
chitin in metal removal was similar enough to chitin to
demonstrate the important role of chitin in these proc-
esses. These data are in agreement with the microbi-
ological data where chitin enhanced microbiological ac-
tivity versus apatite amendments. The chitin amendments
likely stimulated sulfer-transforming bacteria [30,52] and
FeS was observed in the mesocosms. In addition, there
may have been metal sorption to the chitin particles.
However, because chitin is a food source for bacteria,
there is the potential when chitin is digested for metal re-
solubilization. Therefore long-term chitin metal removal
should be evaluated. Nevertheless, the role of apatite
cannot be excluded as phosphate-solubilizing bacteria
were detected. In addition, XAS data also showed the
positive effects of apatite in sediments.
3.4. Effects of Apatite and Chitin
Amendment on Zinc Mobilization
Figure 5 showed an example X-ray absorption near-edge
spectroscopy (XAS) spectra on FAU sediments over time
post amendment addition and reference spectra of pri-
mary components for linear combination fitting (LCF).
Copyright © 2013 SciRes. OJMetal
J. KAN ET AL. 57
Table 3. Metal removal from the overlying water (OW) and
pore-water (PW) samples relative to controls.
Metal removal, %,
OW Metal removal, %, PW
Coarse-grained (YB)
sediment Cu Cr Zn Cu Cr Zn
Apatite (CAS) 40 45 0 NA* NANA
Chitin (CCS) 60 10085 NA NANA
Apatite + chitin (CACS) 70 10057 NA NANA
Fine-grained (MI)
Apatite (FAS) 0 0 24 70 75 45
Chitin (FCS) 62 98 65 87 97 63
Apatite + chitin (FACS) 47 10042 0 68 49
*NA = not analyzed. Pore water was not extractable from these sandy sam-
Figure 5. Normalized Ka Zn X-ray absorption spectra of
FAU sediments over time post amendment addition and
reference spectra of primary components for linear combi-
nation fitting.
Due to concentration of Cu and Cr, as well as detector
issues during data collection, only spectra for Zn were
collected. The coarse samples also provided complica-
tions for data collection due to the low sorption capacity
of sand. Most metals do not sorb to sand, as they tend to
prefer clay minerals and organic matter due to surface
charges. Only treatments FAU, FUS, FAS, FCS and
FACS at certain time points were run on the XAS, and
the Zn XAS linear combination fitting results were
shown in Table 4. Principal component analysis (PCA)
identified and confirmed LCF validity with five suitable
components: Zn hydroxide, Zn Carbonate, Zn orbed to
ferrihydrite, ZnAl2O4, and Zn3(PO4)2 (hopeite).
At day 0, sediments from apatite amended unspiked
treatment (FAU) was identified by XAS to contain pri-
marily easily mobile zinc phases; i.e., zinc hydroxide, Zn
carbonate, Zn sorbed to ferrihydrite, ZnAl2O4, and some
initial Zn3(PO4)2. By day 28 and 120, a significant tran-
sition to sorbed Zn and zinc phosphate was observed in
the presence of the apatite (Table 3). For spiked sedi-
ments, again the predominant initial Zn species is zinc
hydroxide phases (Table 3). Unamended treatment (FUS)
showed no significant changes from easily mobile Zn
hydroxide phase to a more insoluble Zn phosphate. How-
ever, remarkable changes occurred after 28 days post
apatite amendment (FAS) (Table 3), indicating the apa-
tite effectively immobilized and sequestered Zn. Chitin
amendments affected the Zn speciation even further: 1)
~13% hopeite (from zero) in the day 10 formed in chitin
amended sample (FCS) while notable Zn hydroxide
phase decreased from ~63% to 55%; 2) In the apatite +
chitin (FACS), hopeite was observed at day 0 post mix-
ing (~9%), and increased to 27% by Day 10 (Table 3).
Therefore, chitin alone does aid in the transition of solu-
ble zinc to the immobilized phase of hopeite; but the
combined effect of chitin and apatite increased the im-
mobilization by a factor of 2.
4. Conclusion
The apatite plus chitin amendment increased the micro-
bial cell densities and significantly altered the bacterial
population structures (Figures 1 and 4, Table 2). For ex-
ample, growth of Firmicutes was induced, and relative
abundance of Deltaproteobacteria was significantly in-
creased compared to the control (FUS) (Table 2). Del-
taproteobacteria and Firmicutes have both been proven
to be capable of producing sulfides and carbonates that
quickly bind bioavailable metals. Further, phosphate
from the apatite amendments was solubilized biotically
or abiotically. These soluble phosphates sequestered met-
als and formed more thermodynamically stable metal
phosphates. Therefore, we conclude that certain groups
of microorganisms (e.g. Deltaproteobacteria, Firmicute-
sand phosphate-solubilizing bacteria) were induced by
amendments of chitin and apatite, and that these micro-
organisms worked collaboratively and helped facilitate
the transformation and immobilization of the heavy met-
als. Thus, the mixed apatite and chitin amendments may
provide an ideal and cost-effective approach for efficient
and permanent metal sequestration in marine sediments.
These in-situ remedial options show the potential for
more cost effective remediation compared to conven-
tional ex-situ options such as dredging.
5. Acknowledgements
The Strategic Environmental Research and Development
Copyright © 2013 SciRes. OJMetal
Copyright © 2013 SciRes. OJMetal
Table 4. Linear combination fitting of fine-grained sediments (Mare Island Naval Shipyard). Treatments FAU, FUS, FAS,
FCS, and FACS were shown in Table 1. - = not detected. The R factor is the error (%) associated with the fits.
Linear contribution fitting distribution (%)
Treatments Amendment Metal mixture Reaction time
Zn hydroxide1Zn carbonateSorbed Zn2 ZnAl2O4 Hopeite3
R factor
FAU 5% apatite Unspiked 0 days 59.1 7.4 9.5 18.8 5.4 3.00
10 days 59.1 7.9 8.5 18.5 6.2 2.95
28 days 43.4 8.4 18.2 12.9 17.2 1.50
120 days 25.3 4.4 29.9 7.2 33.3 0.35
FUS None Spiked 0 days 71.0 10.0 19.0 - - 1.61
10 days 70.0 10.0 18.0 - - 1.94
28 days 70.0 - 30.0 - -
FAS 5% apatite Spiked 0 days 45.0 16.0 - 14.0 25.0
10 days 43.0 14.0 - 15.0 28.0
28 days 31.0 9.0 26.0 - 34.0
FCS 0.5% chitin Spiked 0 days 63.0 25.0 - 13.0 - 1.12
10 days 55.0 17.0 5.0 11.0 13.0 1.07
FACS 5% apatie + 0.5% chitinSpiked 0 days 68.0 13.0 - 10.0 9.0 1.97
10 days 52.0 15.0 - 8.0 27.0 0.97
1Includes Zn(OH)2 and related layered double hydroxides; 2Zn sorbed to ferrihydrite; 3Zn3(PO4)2.
Program (SERDP) funded this research under project
#ER-1551. The authors thank Gunther Rosen, Ryan Ha-
lonen, Brandon Swope, and Ignacio Rivera for sampling
and technical support. Although EPA contributed to this
article, the research presented was not performed by or
funded by EPA and was not subject to EPA’s quality
system requirements. Consequently, the views, interpre-
tations, and conclusions expressed in this article are
solely those of the authors and do not necessarily reflect
or represent EPA’s views or policies.
[1] R. T. Anderson, H. A. Vrionis, I. Ortiz-Bernad, C. T.
Resch, P. E. Long, R. Dayvault, K. Karp, S. Marutzky, D.
R. Metzler, R. Peacock, D. C. White, M. Lowe and D. R.
Lovley, “Stimulating the in Situ Activity of Geobacter
Species to Remove Uranium from the Groundwater of a
Uranium-Contaminated Aquifer,” Applied and Environ-
mental Microbiology, Vol. 69, No. 10, 2003, pp. 5884-
5891. doi:10.1128/AEM.69.10.5884-5891.2003
[2] B. Sharma, K. H. Gardner, J. Melton, A. Hawkins and G.
Tracey, “Evaluation of Activated Carbon as a Reactive
Cap Sorbent for Sequestration of Polychlorinated Bi-
phenyls in the Presence of Humic Acid,” Environmental
Engineering Science, Vol. 26, No. 9, 2009, pp. 1371-
1379. doi:10.1089/ees.2008.0231
[3] S. Brown, R. Chaney, J. Hallfrisch, J. A. Ryan and W. R.
Berti, “In Situ Soil Treatments to Reduce the Phyto and
Bioavailability of Lead, Zinc, and Cadmium,” Journal of
Environmental Quality, Vol. 33, No. 2, 2004, pp. 522-531.
[4] J. D. Istok, J. M. Senko, L. R. Krumholtz, D. Watson, M.
A. Bogle, A. Peacock, Y. J. Chang and D. C. White, “In
Situ Bioreduction of Technetium and Uranium in a Ni-
trate Contaminated Aquifer,” Environmental Science &
Technology, Vol. 38, No. 2, 2004, pp. 468-475.
[5] T. P. Seager, and K. H. Gardner, “Barriers to Adoption of
Novel Environmental Technologies: Contaminated Sedi-
ments,” In: E. Levner, I. Linkov and J. M. Proth, Eds.,
Strategic Management of Marine Ecosystems, Springer,
2005, pp. 298-312. doi:10.1007/1-4020-3198-X_16
[6] C. J. Werth, R. A. Sanford, R. St John and G. C. Bar-
nuevo, “Long-Term Management of Chlorinated Solvent
Plumes Using a Slow-Release in Situ Electron Donor
Source,” Abstract poster G-4. In: SERDP/ESTCP Meet-
ing, Washington DC, 2005.
[7] H. Paller and A.S. Knox, “Amendments for the in Situ
Remediation of Contaminated Sediments: Evaluation of
Potential Environmental Impacts,” Science of the Total
Environment, Vol. 408, No. 20, 2010, pp. 4894-4900.
[8] Y. J. Chang, P. E. Long, R. Geyer, A. D. Peacock, C. T.
Resch, K. Sublette, S. Pfiffner, A. Smithgall, R. T. An-
derson, H. A. Vrionis, J. R. Stephen, R. Dayvault, I. Ortiz-
Bernad, D. R. Lovley and D. C. White, “Microbial In-
corporation of 13Clabeled Acetate at the Field Scale: De-
tection of Microbes Responsible for Reduction of U(VI),”
Environmental Science & Technology, Vol. 39, No. 23,
2005, pp. 9039-9048. doi:10.1021/es051218u
[9] S. Lukas and J. T. Hollibaugh, “Response of Sediment
Bacterial Assemblages to Selenate and Acetate Amend-
ments,” Environmental Science & Technology, Vol. 35,
No. 3, 2001, pp. 528-534. doi:10.1021/es001492i
J. KAN ET AL. 59
[10] Q. Y. Ma, T. J. Logan and S. J. Traina, “Lead Immobili-
zation from Aqueous Solutions and Contaminated Soils
Using Phosphate Rocks,” Environmental Science & Tech-
nology, Vol. 29, No. 4, 1995, pp. 1118-1126.
[11] S. Knox, D. I. Kaplan and M. H. Paller, “Phosphate
Sources and Their Suitability for Remediation of Con-
taminated Soils,” Science of the Total Environment, Vol.
357, No. 1-3, 2006, pp. 271-279.
[12] X. D. Cao, L. Q. Ma and A. Wahbi, “Immobilization of
Cu, Zn, and Pb in Contaminated Soils Using Phosphate
Rock and Phosphoric Acid,” Journal of Hazardous Mate-
rials, Vol. 164, No. 2-3, 2009, pp. 555-564.
[13] Q. Y. Ma, S. Traina, T. Logan and J. Ryan, “In Situ Lead
Immobilization by Apatite,” Environmental Science &
Technology, Vol. 27, No. 9, 1993, pp. 1803-1810.
[14] J. Wright, K. R. Rice, B. Murphy and J. Conca, “PIMS
Using Apatite IITM: How It Works To Remediate Soil
and Water,” In: R. E. Hinchee and B. Alleman, Eds., Sus-
tainable Range Management, Battelle Press, Columbus,
[15] U. Ghosh, R. G. Luthy, G. Cornelissen, D. Werner and C.
A. Menzie, “In-Situ Sorbent Amendments: A New Direc-
tion in Contaminated Sediment Management,” Environ-
mental Science & Technology, Vol. 45, No. 4, 2011, pp.
1163-1168. doi:10.1021/es102694h
[16] G. B. Williams, K. G. Scheckel, G. McDermott, D. Grat-
son, D. Neptune and J. A. Ryan, “Speciation and Bio-
availability of Zinc in Amended Sediments,” Chemical
Speciation and Bioavailability, Vol. 23, No. 3, 2011, pp.
143-154. doi:10.3184/095422911X13103699236851
[17] B. Uzair and N. Ahmed, “Solubilization of Insoluble In-
organic Phosphate Compounds by Attached and Free-
Living Marine Bacteria,” Journal of Basic & Applied Sci-
ences, Vol. 3, No. 2, 2007, pp. 59-63.
[18] C. Kanzog, A. Ramette, N. V. Queric and M. Klages,
“Response of Benthic Microbial Communities to Chitin
Enrichment: An in Situ Study in the Deep Arctic Ocean,”
Polar Biology, Vol. 32, No. 1, 2008, pp. 105-112.
[19] A. Boetius and K. Lochte, “Effect of Organic Enrich-
ments on Hydrolytic Potentials and Growth of Bacteria in
Deep-Sea Sediments,” Marine Ecology Progress Series,
Vol. 140, 1996, pp. 239-250. doi:10.3354/meps140239
[20] B. L. Bassler, C. Yu, Y. C. Lee and S. Roseman, “Chitin
Utilization by Marine Bacteria: Degradation and Catabo-
lism of Chitin Oligosaccharides by Vibrio furnissii,” Jour-
nal of Biological Chemistry, Vol. 266, No. 36, 1991, pp.
[21] L. Brierley, “Metal Immobilization Using Bacteria,” In: H.
L. Ehrlich and C. L. Brierley, Eds., Microbial Mineral
Recovery, McGraw-Hill, 1990, pp. 303-323.
[22] B. M. Tebo, “Metal Precipitation by Marine Bacteria:
Potential for Biotechnological Applications,” In: J. K.
Setlow, Ed., Genetic Engineering, Plenum Press, 1995,
pp. 231-261.
[23] R. Lovley, “Dissimilatory Metal Reduction,” Annual Re-
view of Microbiology, Vol. 47, 1993, pp. 263-290.
[24] L. J. Barnes, P. J. M. Scheeren and C. J. N. Buisman,
“Microbial Removal of Heavy Metal and Sulfate from
Contaminated Groundwaters,” In: J. L. Means and R. E.
Hinchee, Eds., Emerging Technology for Bioremediation
of Metals, CRC Press, 1994, pp. 38-49.
[25] L. L. Barton and F. A. Tomei, “Characteristics and Ac-
tivities of Sulfate-Reducing Bacteria,” In: L. L. Barton,
Ed., Sulfate-Reducing Bacteria, Plenum Press, 1995, pp.
1-32. doi:10.1007/978-1-4899-1582-5_1
[26] R. T. Anderson and D. R. Lovley, “Ecology and Biogeo-
chemistry of in Situ Groundwater Bioremediation,” In: J.
G. Jones, Ed., Advances in Microbial Ecology, Plenum
Press, 1997, pp. 289-350.
[27] K. H. Nealson, “Sediment Bacteria: Who’s There, What
Are They Doing, and What’s New?” Annual Review of
Earth and Planetary Sciences, Vol. 25, 1997, pp. 403-434.
[28] B. M. Tebo and A. Y. Obraztsova, “Chromium(VI), Man-
ganese(IV), Uranium(VI), and Iron(III): Electron Accep-
tors for Growth for a Novel Spore Forming Sulfate Re-
ducing Bacterium,” FEMS Microbiology Letters, Vol.
162, No. 1, 1998, pp. 193-198.
[29] Y. M. Arias and B. M. Tebo, “Comparative Studies of
Cr(VI) Reduction by Sulfidogenic and Non-Sulfidogenic
Microbial Communities,” Applied and Environmental Mi-
crobiology, Vol. 69, No. 3, 2003, pp. 1847-1853.
[30] J. Kan, Y. Wang, A. Obraztsova, G. Rosen, J. Leather, K.
G. Scheckel, K. H. Nealson and Y. M. Arias-Thode, “Ma-
rine Microbial Community Response to Inorganic and Or-
ganic Sediment Amendments in Laboratory Mesocosms,”
Ecotoxicology and Environmental Safety, Vol. 74, No. 7,
2011, pp. 1931-1941. doi:10.1016/j.ecoenv.2011.06.011
[31] G. Rosen, J. Leather, J. Kan and Y. M. Arias-Thode,
“Ecotoxicological Response of Marine Organisms to In-
organic and Organic Sediment Amendments in Labora-
tory Exposures,” Ecotoxicology and Environmental Safety,
Vol. 74, No. 7, 2011, pp. 1921-1930.
[32] T. Huber, G. Faulkner and P. Hugenholtz, “Bellerophon;
A Program to Detect Chimeric Sequences in Multiple
Sequence Alignments,” Bioinformatics, Vol. 20, No. 14,
2004, pp. 2317-2319. doi:10.1093/bioinformatics/bth226
[33] K. Ayyakkannu and D. Chandramohan, “Occurrence and
Distribution of Phosphate Solubilizing Bacteria and Phos-
phatase in Marine Sediments at Porto Novo,” Marine Bi-
ology, Vol. 11, No. 3, 1971, pp. 201-205.
[34] J. Talbot and A. Weiss, “Laboratory Methods for ICP-MS
Analysis of Trace Metals in Precipitation,” 1994.
[35] S. E. Bufflap and H. E. Allen, “Comparison of Pore Wa-
ter Sampling Techniques for Trace Metals,” Water Re-
search, Vol. 29, No. 9, 1995, pp. 2051-2054.
Copyright © 2013 SciRes. OJMetal
Copyright © 2013 SciRes. OJMetal
[36] G. Scheckel, J. A. Ryan, D. Allen and N. V. Lescano,
“Determining Speciation of Pb in Phosphate-Amended
Soils: Method Limitations,” Science of the Total Envi-
ronment, Vol. 350, No. 1-3, 2005, pp. 261-272.
[37] H. M. Cauchie, “Chitin Production by Arthropods in the
Hydrosphere,” Hydrobiology, Vol. 470, No. 1-3, 2002, pp.
63-96. doi:10.1023/A:1015615819301
[38] M. Poulicek and C. Jeuniaux, “Chitin Biodegradation in
Marine Environments: An Experimental Approach,” Bio-
chemical Systematics and Ecology, Vol. 19, No. 5, 1991,
pp. 385-394. doi:10.1016/0305-1978(91)90055-5
[39] J. W. Deming and J. A. Baross, “The Early Diagenesis of
Organic Matter: Bacterial Activity,” In: M. H. Engel and
S. A. Macko, Eds., Organic Geochemistry: Principles
and Application, Plenum Press, New York, 1993, pp.
119-144. doi:10.1007/978-1-4615-2890-6_5
[40] A. Bell, J. C. Hubbard, L. Liu, R. M. Davis and K. V.
Subbarao, “Effects of Chitin and Chitosan on the Inci-
dence and Severity of Fusarium Yellows in Celery,”
Plant Disease, Vol. 82, No. 3, 1998, pp. 322-328.
[41] J. Hallmann, R. Rodriguez-Kabana and J. W. Kloepper,
“Chitin-Mediated Changes in Bacterial Communities of
the Soil, Rhizosphere and within Roots of Cotton in Rela-
tion to Nematode Control,” Soil Biology and Biochemis-
try, Vol. 31, No. 4, 1999, pp. 551-560.
[42] D. M. Akob, H. J. Mills and J. E. Kostka, “Metabolically
Active Microbial Communities in Uranium-Contaminated
Subsurface Sediment,” FEMS Microbiology Ecology, Vol.
59, No. 1, 2006, pp. 95-107.
[43] G. Garau, P. Castaldi, L. Santona, P. Deiana and P. Melis,
“Influence of Red Mud, Zeolite and Lime on Heavy
Metal Immobilization, Culturable Heterotrophic Micro-
bial Populations and Enzyme Activities in a Contami-
nated Soil,” Geoderma, Vol. 142, No. 1-2, 2007, pp. 47-
57. doi:10.1016/j.geoderma.2007.07.011
[44] J. Kleikemper, O. Pelz, M. H. Schroth and J. Zeyer, “Sul-
fate-Reducing Bacterial Community Response to Carbon
Source Amendments in Contaminated Aqufer Micro-
cosms,” FEMS Microbiology Ecology, Vol. 42, No. 1,
2002, pp. 109-118.
[45] L. L. Barton and G. D. Fauque, “Biochemistry, Physiol-
ogy and Biotechnology of Sulfate-Reducing Bacteria,”
Advances in Applied Microbiology, Vol. 68, 2009, pp.
41-98. doi:10.1016/S0065-2164(09)01202-7
[46] C. Myers and K. H. Nealson, “Bacterial Manganese Re-
duction and Growth with Manganese Oxide as the Sole
Electron Acceptor,” Science, Vol. 240, No. 4857, 1988,
pp. 1319-1321. doi:10.1126/science.240.4857.1319
[47] G. N. Baturin, “Some Unique Sedimentological and Geo-
chemical Features of Deposits in Coastal Upwelling Re-
gions,” In: J. Thiede and E. Suess, Eds., Coastal Upwell-
ing, Plenum Press, 1983, pp. 11-27.
[48] C. E. Reimers, M. Kastner and R. E. Garrison, “The Role
of Bacterial Mats in Phosphate Mineralization with Par-
ticular Reference to the Monterey Formation,” In: W. C.
Burnett and S. R. Riggs, Eds., Phosphate Deposits of the
World, Cambridge University Press, 1990, pp. 300-311.
[49] N. Schulz and H. D. Schulz, “Large Sulfur Bacteria and
the Formation of Phosphorite,” Science, Vol. 307, 2005,
pp. 416-418. doi:10.1126/science.1103096
[50] H. Rodriguez and R. Fraga, “Review: Phosphate Solubi-
lizing Bacteria and Their Role in Plant Growth Promo-
tion,” Biotechnology Advances, Vol. 17, No. 4-5, 1999,
pp. 319-339. doi:10.1016/S0734-9750(99)00014-2
[51] D. De Souza, S. Nair and D. Chandramohan, “Phosphate
Solubilizing Bacteria around Indian Peninsula,” Indian
Journal of Marine Sciences, Vol. 29, 2000, pp. 48-51.
[52] A. Robinson-Lora and R. A. Brennan, “Efficient Metal
Removal and Neutralization of Acid Mine Drainage by
Crab-Shell Chitin under Batch and Continuous Flow Con-
ditions,” Bioresource Technology, Vol. 100, No. 21, 2009,
pp. 5063-5071. doi:10.1016/j.biortech.2008.11.063
J. KAN ET AL. 61
(a) (b)
Figure S1. DGGE fingerprints of bacterial community from aerobic/anaerobic interface (a) and anaerobic zone (b) sedi-
ments. Labels referred to the corresponding treatments as listed in Table 1. Bands 16-29 were selected and excised for se-
quencing. Anaerobic sediment samples from treatments FCS and FACS were not available (b).
haeum equitans AJ318041
DGGE band 21
Uncultured Spirochaeta sp. FJ752792
Uncultured bacterium GQ853685
Uncultured Spirochaeta sp. AB121108
Roseobacter sp. AY576690
DGGE band 22
Roseobacter sp. EU195946
DGGE band 24
Uncultured gammaproteobacterium FN397816
Endosymbiont bacterium AJ441188
Endosymbiont bacterium AJ441189
DGGE band 16
Uncultured deltaproteobacterium FJ753016
Uncultured bacterium GQ246304
Uncultured deltaproteobacterium AJ969448
Delsulfotalea sp. AJ318381
Uncultured deltaproteobacterium AF432273
DGGE band 18
Uncultured bacterium EU142058
Uncultured deltaproteobacterium FJ664778
Uncultured deltaproteobacterium FJ664813
DGGE band 27
DGGE band 17
DGGE band 28
DGGE band 19
Cyclobacterium sp. AY349464
Cytophaga sp. AM180747
Cytophaga sp. EU768822
Uncultured bacterium FM201245
DGGE band 23
Uncultured Bacteroidetes AY263723
DGGE band 25
Uncultured Bacteroidetes GU061313
Uncultured bacterium AY171354
Uncultured bacterium DQ334660
DGGE band 20
Uncultured bacterium DQ3346601
DGGE band 26
Uncultured bacterium FJ716908
Uncultured Bacteroidetes EU035609
Uncultured Bacteroidetes EU035608
DGGE band 29
Uncultured bacterium GQ342175
Uncultured bacterium EU617867
Uncultured bacterium GU066435
Figure S2. Phylogenetic analysis (DNA distance—NJ) of
bacterial DGGE band sequences obtained from Supplemen-
tary Figure 2. Alpha, Gamma, and Delta represent subdivi-
sions of Proteobacteria. Bootstrap values were based on
1000 replicated trees. Nanoarchaeumequitans was used as
an outgroup. Scale bar represent 0.05 substitutions per site.
Figure S3. PSB (phosphate solubilizing bacteria) cultures
after 2 weeks incubation. Clearing zones showed bacterial
solubilization of calcium phosphate. Left: FAS treatment
with 104 dilution; Right: FACS treatment with 104 dilu-
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