functional diversity changes of microbial communities along a soil aquifer for reclaimed water...
TRANSCRIPT
R E S EA RCH AR T I C L E
Functional diversity changes of microbial communities along asoil aquifer for reclaimed water recharge
Xue Zhang, Xuan Zhao & Meng Zhang
Laboratory of Environmental Technology, Institute of Nuclear and New Energy Technology, Tsinghua University, Beijing, China
Correspondence: Xuan Zhao, Laboratory of
Environmental Technology, Institute of
Nuclear and New Energy Technology,
Tsinghua University, Beijing 100084, China.
Tel.: +8610 62796428; fax: +8610
62771150; e-mail: [email protected]
Received 12 July 2011; revised 15 October
2011; accepted 14 November 2011.
Final version published online 11 January
2012.
DOI: 10.1111/j.1574-6941.2011.01263.x
Editor: Julian Marchesi
Keywords
groundwater recharge; soil aquifer
treatment; BIOLOG assay; soil-attached
microbial community; reclaimed water.
Abstract
The physiochemical and functional diversity of soil-attached microorganisms
was investigated using a stabilized laboratory-scale soil aquifer treatment (SAT)
system. In this system, reclaimed water after ozonation was used as the feed
water, and 60% dissolved organic carbon was removed by the unsaturated
vadose layer in 0.8 days. Soil biomass (volatile solids, phospholipid extraction)
and functional diversity significantly decreased from the unsaturated vadose
layer to the saturated aquifer, where they maintained the same level. Using
principal components analysis based on substrate utilization pattern, the vadose
layer soil sample was clearly separated from the saturated layer samples. Excep-
tionally, the oxidation rates of esters remained stable during SAT, indicating
the purification potential on certain recalcitrant organic compounds in the sat-
urated aquifer given an adequate retention time. Correlation analysis revealed
that organic carbon was the key limiting factor for microbial biomass and
activity, especially for tyrosine-like aromatic proteins and soluble microbial
byproduct-like materials.
Introduction
Water stress attributable to water scarcity and quality deg-
radation is occurring in many regions of the world.
Reclaimed water, considered a promising alternative water
resource, has as a result gained an increasing level of atten-
tion. Artificial groundwater recharge (AGR) used with
reclaimed water is an attractive option for water reuse
because of the additional advantages in seasonal and long-
term water storage and control of saltwater intrusion
(Miller, 2006). During AGR, soil aquifer treatment (SAT)
provides final purification of the reclaimed water and is
considered a sustainable, effective and economic method of
organic matter removal (Fox et al., 2005; Zhao et al.,
2009). In SAT, removal of organic matter is primarily
attributed to biodegradation, especially aerobic biodegrada-
tion (Quanrud et al., 2003; Kolehmainen et al., 2007; Xue
et al., 2009). Thus, the microbial community in SAT plays
an important role in the attenuation of organic pollutants.
The dynamics of microbial communities and their role
in pollutant removal is one of the hottest topics in SAT.
A large number of studies on microorganisms suspended
in pore water have revealed that during SAT, the bacterial
cell concentration/extracellular enzyme activities (EEAs),
e.g. a-D-glucosidase and phosphomonoesterase, were sig-
nificantly correlated with nutrient concentrations [e.g.
dissolved organic carbon (DOC)/biodegradable dissolved
organic carbon (BDOC)] of the water (Hendel et al.,
2001; Kolehmainen et al., 2007, 2009). Moreover, the bac-
terial community changed from an Actinobacteria-domin-
anted population in lake water to a diverse and then
primary proteobacterial community after travelling 0.6 m
in the sand column (Kolehmainen et al., 2008).
However, the presence of soil-attached microbial bio-
masses cannot be disregarded. Numerous studies on
aquatic ecosystems have shown higher cell-specific activi-
ties of attached vs. suspended bacteria and have provided
evidence that the majority of biodegradation occurs
through soil-attached microbial communities (Harvey
et al., 1984; Middelboe et al., 1995; Grossart & Simon,
1998). Some indicators, such as EEAs and soil biomass,
have been used to characterize the dynamics of soil
microbial communities during SAT. A strong positive
correlation was found between total viable soil biomass
and organic carbon removal in SAT (Rauch & Drewes,
2005; Rauch-Williams & Drewes, 2006). In using EEAs,
FEMS Microbiol Ecol 80 (2012) 9–18 ª 2011 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
MIC
ROBI
OLO
GY
EC
OLO
GY
no major differences were discerned between the surface
and bottom sediments (Kolehmainen et al., 2009). How-
ever, Schutz et al. (2010) found a significant decrease in
EEAs with soil depth. Different results may be attribut-
able to different sites or to the relatively low sensitivity of
the methods. Although soil biomass and enzyme activities
provide some data, the dynamics and the role of the
microbial community during SAT remain relatively
unknown. Novel sensitive methods need to be applied to
characterize microbial communities.
Generally, functional diversity is essential in understand-
ing the role of microbial communities in different environ-
ments (Preston-Mafham et al., 2002). The BIOLOG assay
has proved to be a useful and sensitive method in distin-
guishing functional differences among microbial commu-
nities from various habitats (Garland & Mills, 1991;
Firestone et al., 1998; Weber et al., 2008; Al-Mutairi,
2009). Therefore, the BIOLOG assay is helpful in the inter-
pretation of biodegradation capabilities during SAT.
In the present study, the BIOLOG assay is used to
distinguish the functional diversity of soil-attached micro-
bial communities in laboratory-simulated SAT systems,
wherein secondary effluent after ozonation was supplied as
feed water. The appearance, distributions and biomasses of
soil-attached microbial communities were detected using
environmental scanning electron microscopy (ESEM),
analysis of volatile solids (VS) and phospholipid extraction
(PLE). Thereafter, correlations between the characteristics
of the microbial communities (biomass, functional diver-
sity, metabolic activity) and the nutrient availability of feed
water were analysed to identify the role of microbes in SAT.
Materials and methods
The laboratory-scale SAT system
The SAT system was simulated using five soil columns
(Fig. 1). The internal diameters were 12 cm for the first
column (C1) and 24 cm for the other four columns
(C2–C5). All five columns were 200 cm in height, with a
packed-bed height of 180 cm. These columns were filled
with sandy powdery soil (grain size 0.4–0.8 mm) with
porosity of 0.39 collected from a 9- to 17-m-deep aquifer
in a suburb of Beijing. The C1 column was operated
under unsaturated conditions by pumping reclaimed
water into the top of the column, with the cycle compris-
ing 3 days of flooding and 1 day of drying. This column
was used to simulate the vadose soil layer. The other four
columns had sealed lids and were operated under satu-
rated conditions (constant flooding), representing the
saturated aquifer layer. All columns were kept at room
temperature (20 ± 2 °C) in the dark.
The secondary effluent from the Gaobeidian Wastewa-
ter Treatment Plant (WWTP), which used a traditional
activated sludge treatment process, was supplied as the
feed water of C1 after the addition of 5 mg L�1 ozone.
The ozone dosage was optimized in a previous study to
achieve a higher BDOC/DOC ratio and enhanced biode-
gradability (Liu, 2003). The effluent of C1 was stored in a
10-L tank, and a portion of this effluent was pumped in
sequence through columns C2–C5. The cumulative
hydraulic retention times of columns C1–C5 were 0.8,
7.8, 14.8, 21.8 and 28.8 days, respectively. The system was
biologically adapted and ready for sampling approxi-
mately after 1 year of acclimatization, with stable quality
effluents monitored at least once a week.
Water sample collection and physiochemical
analysis
The secondary effluent from the Gaobeidian WWTP (R0)
and the effluents after ozonation (R1), as well as all
the effluents from the bottom of the five soil columns
(C1–C5) were identified as WS1, WS2, WS3, WS4 and
WS5, respectively, and were sampled at least once a week.
All water samples were first filtered using a 0.45-lm filer,
and the following parameters were then measured:
DOC, absorbance at 254 nm (UV254), nitrate (NO3-N),
ammonia (NH4-N), phosphate (PO4-P) and three-dimen-
sional excitation-emission matrix (EEM) spectra. DOC
Tank
C1
2
C2 C3 C5 C4
Ozone detector
Ozonizor
Ozone decomposer
Tank for R0 Ozone reactor
Fig. 1. Schematic diagram of the laboratory-
scale AGR system.
ª 2011 Federation of European Microbiological Societies FEMS Microbiol Ecol 80 (2012) 9–18Published by Blackwell Publishing Ltd. All rights reserved
10 X. Zhang et al.
was measured using a Shimadzu Total Organic Carbon
Analyzer (TOC-VWP, Shimadzu Corp., Japan). UV254 was
analysed using a Shimadzu UV-3100 UV-visible spectro-
photometer (Shimadzu) at 254 nm. Nitrate (NO3-N),
ammonia (NH4-N) and phosphate (PO4-P) were measured
according to the specifications of the Ministry of Environ-
mental Protection PR China (2002). The average water
quality parameters of 5 weeks’ worth of records (2 weeks
prior, 2 weeks after and the week during soil sampling) are
listed in the Results. EEM spectra were measured using an
F-7000 FL spectrophotometer (Hitachi, Japan) to identify
the dissolved organic matter (DOM) composition changes.
The EEM spectra were a collection of corresponding scan-
ning emission spectra (Em) from 280 to 550 nm at 2-nm
increments through variation of the excitation wavelength
(Ex) from 220 to 450 nm at 5-nm sampling intervals. The
excitation and emission slits were maintained at 5 nm, and
the scanning speed was set at 1200 nm min�1. The spec-
trum of super-Q water was recorded as the blank control
and subtracted from the EEM spectra of all samples. The
normalized region-specific excitation–emission area vol-
ume (ui,n) and the percentage fluorescence response (Pi,n)
of region i were calculated using the fluorescence regional
integration (FRI) technique (Chen et al., 2003).
Soil sample collection and preparation
After 1 year of operation, the physicochemical parameters
of feed water and all effluents in the SAT were stable,
implying that mature and steady microbial communities
were formed during SAT. Four soil samples (equal to
50 g dry soil weight each) were scratched from the top
(depth of 5–10 cm) of columns C1–C4 using a clean steel
scoop. The four soil samples were identified as S1, S2, S3
and S4, respectively, representing soil samples from differ-
ent soil depths. All samples were stored in sterile blue-
capped bottles at 4 °C before analysis.
ESEM examination
Micrographs of the four soil samples were obtained using
ESEM (FEI Quanta 200; FEI, Czech Republic) without
any pretreatments. One clean sand sample prepared from
1 mL S4 and was used as the blank control. This control
sample was first washed with 1% NaOH to remove
microorganisms before being observed by ESEM.
Soil biomass quantification
Total biomass accumulation in the soil was identified as
VS and phospholipids. VS were identified according to
standard APHA 2540 (2005) through sand combustion at
550 °C. PLE quantifies the amount of phospholipids in
the cell walls of viable cells biomass using a colour reac-
tion with ammonium molybdate and malachite green, as
described by Rauch-Williams & Drewes (2006). Each soil
sample was measured in triplicate.
BIOLOG assay
The microbial community was first extracted from the
soil samples. Each soil sample (equal to 45 g dry weight)
was suspended in 35 mL sterile saline solution (0.85%
NaCl) on a rotary shaker at 240 r.p.m. for 30 min at
25 °C and then shaken in an ultrasonic cleaning machine
(Shanghai Lvyu Biotech Co., KQ-500E, 40 kHz, 100 W)
for 3 min. The microorganisms were proven to be effec-
tively detached from the soil without lysis during the pro-
cesses (Joyce et al., 2003). The suspensions were allowed
to settle for 5 min. The supernatants were transferred to
50-mL tubes and centrifuged at 1400 g for 5 min. The
supernatants were again transferred to new tubes and
diluted with sterile saline solution, resulting in a uniform
OD600 nm of 0.06 cm�1 (Wang et al., 2009). Microbial
suspensions of 150 lL were added to each well of the
BIOLOG microplate, and were incubated under aerobic
conditions at 30 °C in the dark.
Each BIOLOG ECO plate (Biolog Inc., Hayward, CA)
consists of three replicates of 96 wells, each comprising
31 sole carbon sources and one water blank control.
Metabolism of the substrate in particular wells results in
a colour change in the tetrazolium dye. Absorbance read-
ings of the plates were taken at 595 nm every 4 h for
168 h, using a microtitre plate reader (MD SpectraMax
MS, MD Inc.).
Data analysis
The well absorbance values of the BIOLOG plate were
adjusted through subtraction of the blank control well.
The average well colour development (AWCD), calculated
as the average adjusted absorbance of all wells per plate,
was used as an indicator of general microbial activity
(Garland & Mills, 1991). The absorbance data for the 31
individual carbon substrates at 72.6 h was used to quan-
tify C-source utilization using principal components anal-
ysis (PCA) because the AWCD value at 72.6 h preserved
the greatest variance between well responses while keeping
the maximum number of wells within the linear absor-
bance range (Weber et al., 2008). PCA was performed
using SPSS software with standardized absorbance data
(Wang et al., 2009). Differences in microbial functional
diversity among soil samples were also compared using the
mean Shannon–Weaver indices. Following the formula of
Al-Mutairi (2009), richness (R, the number of oxidized
carbon substrates) and the Shannon–Weaver index (i.e.
FEMS Microbiol Ecol 80 (2012) 9–18 ª 2011 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
Microbial dynamic during SAT for water recharge 11
diversity H′ and evenness index J′) were calculated using
an OD of 0.25 as the threshold for positive response (Go-
mez et al., 2006). The substrate oxidation rate (cm�1 h�1)
was calculated as the slope of the colour development
curve (e.g. AWCD curve) during its linear phase.
Pearson product-moment correlations among the
parameters were conducted using SPSS 14.0 software (SPSS
Inc.).
Results and discussion
Physiochemical characteristics of water during
the laboratory-scale SAT
The physicochemical parameters of water quality at each
sampling point are listed in Table 1. The organic matter
content was presented as DOC, UV254 and SUVA (spe-
cific UVA, which is equal to UV254/DOC). Robust
removal of DOC in laboratory-scale SAT was certified
during the 1 year of operation. DOC did not decrease
after ozonation, whereas more than 60% of the DOC was
removed within a 1.8-m unsaturated vadose soil layer
(column C1), and an additional 19% was removed after
7 days travel within the first 1.8 m of saturated aquifer
treatment (column C2). These results are consistent with
previous laboratory and field studies (Quanrud et al.,
2003; Vanderzalm et al., 2006; Zhao et al., 2009). The
long-term operation of SAT with high and constant
removal efficiency of organic pollutants indicates that
biodegradation, rather than adsorption, is the primary
process, as has also been shown using abiotic control in
previous studies (Quanrud et al., 2003). Based on the
high removal efficiency of DOC by the unsaturated
vadose layer, aerobic degradation was considered to be
primarily responsible for DOC removal.
UV254 decreased by 60% after ozonation, whereas 13%
and 10% removal was achieved along columns C1 and
C2, respectively. However, both DOC and UV254 changed
slightly in later saturated aquifer treatments (columns C3
–C5). SUVA, which represents the relative aromaticity of
the bulk organic compounds, decreased dramatically after
ozonation, but increased during SAT. These results indi-
cated that aromatic compounds are preferentially
destroyed by ozonation, whereas more aliphatic com-
pounds are removed by the sand columns. The increase
in SUVA during SAT may be attributable to enrichment
of aromatic compounds through biodegradation (Xue
et al., 2009).
Most NH4-N was removed during ozonation in the
unsaturated vadose layer, whereas slight denitrification
occurred in the saturated aquifer, and total nitrogen
could be reduced by 1.5–2.0 mg L�1. Most PO4-P was
removed in the first 7 days of travel in the saturated aqui-
fer (column C2). SAT ran steadily with good purification
performance for over 1 year, indicating its stability.
ESEM micrographs and biomass qualification of
the soil samples
ESEM micrographs of sand samples S1–S4 were obtained
at various magnifications (Fig. 2). Obvious biofilms were
found on the surface of S1 and S2, where bacteria sur-
rounded by extracellular mucilages appeared to be the
primary colonizers. The appearance of S3 and S4 was sig-
nificantly similar to clean sand (blank control), indicating
the existence of fewer microorganisms in the deeper satu-
rated aquifer after 7 days of travel. Some unicellular
eukaryotes (algae/fungi/prozotoa) were also found in sand
samples S1 and S2, but were sparse in samples S3 and S4.
Based on the ESEM micrographs, an obvious decrease
in biofilm formation was found during SAT. The biomass
contents of soils were determined quantitatively as VS
and phospholipids. The VS content, representing the total
biomass, was clearly higher in S1 (7.8 mg g�1) than in
the other three samples, whereas S2, S3 and S4 had simi-
lar VS contents, ranging from 5.3 to 5.8 mg g�1
(Table 2). Similar results were determined based on PLE,
the phospholipids representing the viable biomass. The
phospholipid content in S1 (32.6 nmol PO3�4 g�1) was
significantly higher than in the other three soil samples,
Table 1. Physicochemical water quality at each sampling point
Sample R0* R1* WS1* WS2* WS3* WS4* WS5*
DOC (mg L�1) 4.35 (0.25) 4.31 (0.30) 1.73 (0.62) 0.91 (0.24) 0.87 (0.07) 0.82 (0.10) 0.91 (0.14)
UV254 (m�1) 11.50 (0.34) 4.44 (1.39) 2.92 (1.06) 1.84 (0.44) 1.88 (0.29) 1.72 (0.33) 2.26 (0.21)
SUVA [L (mg m)�1] 2.64 1.03 1.69 2.03 2.15 2.11 2.49
NH4-N (mg L�1) 1.02 (0.75) 1.08 (0.69) 0.18 (0.03) 0.14 (0.02) 0.13 (0.01) 0.13 (0.02) 0.13 (0.01)
NO3-N (mg L�1) 21.92 (4.44) 21.88 (4.23) 23.17 (2.84) 22.86 (1.89) 20.73 (0.98) 20.61 (0.71) 20.88 (0.76)
PO4-P (mg L�1) 2.49 (0.35) 2.43 (0.35) 1.68 (0.99) 0.73 (0.01) 0.91 (0.05) 1.09 (0.10) 0.88 (0.03)
pH 7.27 (0.21) 7.34 (0.18) 7.64 (0.31) 7.94 (0.28) 7.70 (0.30) 7.73 (0.26) 8.03 (0.24)
Data are presented as the mean (SD) of 5-week records.
*Water samples: R0, secondary effluent from Gaobeidian WWTP; R1, water sample after ozonation; WS1, WS2, WS3, WS4 and WS5, effluents
obtained from bottom of the five soil columns (C1–C5), respectively.
ª 2011 Federation of European Microbiological Societies FEMS Microbiol Ecol 80 (2012) 9–18Published by Blackwell Publishing Ltd. All rights reserved
12 X. Zhang et al.
Fig. 2. ESEM micrographs of the sand from
the top of different sand columns and one
washed sand sample (S1–S4).
Table 2. Soil biomass and microbial functional diversity of the four soil samples*
Sample VS (mg g�1)†PLE (nmol
PO3�4 g�1)† H′ R J′
S1** 7.5 (0.8) 32.6 (3.8) 2.97 (0.05) 30 (1.1) 0.87 (0.01)
S2** 5.8 (0.6) 13.0 (2.6) 0.95 (0.46) 8 (4.0) 0.45 (0.10)
S3** 5.3 (0.9) 13.3 (5.1) 1.04 (0.26) 6 (2.1) 0.58 (0.04)
S4** 5.7 (0.7) 13.9 (4.7) 1.78 (0.02) 11 (0) 0.74 (0.01)
*Data are presented as the mean (SD) of triplicate experiments. H′, Shannon–Weaver diversity index; R, number of oxidized carbon substrates; J′,
Shannon–Weaver evenness index.
**S1, S2, S3, S4 were four soil samples taken from the top of columns C1–C4, respectively.†Given as the amount per gram of dry soil.
FEMS Microbiol Ecol 80 (2012) 9–18 ª 2011 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
Microbial dynamic during SAT for water recharge 13
which ranged from 13.0 to 13.9 nmol PO3�4 g�1. Both VS
and PLE data indicate that biomass in the soil sharply
decreased from the unsaturated vadose soil layer to the
saturated aquifer layer, whereas biomass remained at the
same level in the saturated layer (S2–S4). The decrease in
soil biomass co-occurred with the decrease in DOC
concentration during SAT.
BIOLOG assay of the soil samples
Using BIOLOG Eco plates, the metabolic activities of the
soil-attached microbial community were studied during
SAT. AWCD, as a measure of total microbial metabolic
activity, generally followed similar patterns with incuba-
tion time (Fig. 3). Clearly, a higher AWCD value was
detected in sample S1 than in the other three soil sam-
ples. The average substrate oxidation rate of S1 was
0.025 cm�1 h�1, approximately four times higher than
those of the other samples (0.005–0.007 cm�1 h�1). These
results indicate that significantly higher metabolic activity
was detected in the microbial community associated with
unsaturated vadose soil. Moreover, the lag time prior to
colour development was significantly shorter for S1
(25 h) than for the other three soil samples (over 50 h).
The lag time is determined by the initial inoculum den-
sity and relative growth rates of the species that are capa-
ble of utilizing the carbon source within each well
(Preston-Mafham et al., 2002). The four suspensions were
incubated at the same cell densities on the BIOLOG
plates, so the variance in lag time resulted from the dif-
ferent growth rates of the microbial communities. Thus,
the microbial community in sample S1 actively respired
and grew faster than those in the other samples. The
obviously higher metabolic activity in the top unsaturated
soil layer can explain the bulk organic carbon removal
within the upper vadose layer of 1.0–2.0 m (Drewes &
Fox, 1999; Rauch-Williams & Drewes, 2006; Vanderzalm
et al., 2006).
More specifically, the carbon sources in the BIOLOG
Eco plate were divided into seven groups (Jena et al.,
2006). A comparison of the substrate oxidation rates of
individual carbon source groups was then conducted
(Fig. 4). Most of the substrate (except for esters) oxida-
tion rates in sample S1 were significantly higher (approxi-
mately 2.3–9.5 times higher) than those in S2–S4,whereas those in the latter three (S2–S4) samples were at
roughly the same level. These results are consistent with
the average metabolic rates, which revealed a significant
decrease in metabolic activities from the unsaturated
column to the saturated aquifer. Exceptionally, the
oxidation rates of esters were all approximately 0.020
–0.028 cm�1 h�1, revealing their uniform metabolic capa-
bilities during SAT. Additionally, the lag times for ester
oxidation were 22 and 31 h for S1 and S2–S4, respec-
tively. The differences in lag time were significantly smal-
ler for esters than for AWCD. The ester-degrading
microorganisms were distributed evenly and remained
active during SAT, in dramatic contrast to other microor-
ganisms.
Regarding the unique distribution of ester-degrading
bacteria, there are two aspects to consider. First, the con-
tinuous presence of certain carbon sources is required in
the recharged water, meaning that such carbon sources
are refractory. Humic acids, fluvic acids and certain trace
pollutants (i.e. sulfamethoxazole and EDTA) are reported
as refractory compounds, possibly producing these results
(Drewes & Jekel, 1998; Conn et al., 2010; Maeng et al.,
Fig. 3. AWCD of the BIOLOG Eco plates for microbial samples from
different sand columns.
Fig. 4. Substrate oxidation rate of each carbon source group with
different microbial communities (CH, carbohydrates; CA, carboxylic
acids; AA, amino acids; PC, phosphorylated compounds).
ª 2011 Federation of European Microbiological Societies FEMS Microbiol Ecol 80 (2012) 9–18Published by Blackwell Publishing Ltd. All rights reserved
14 X. Zhang et al.
2011). Secondly, the uniform metabolic activity of certain
microorganisms indicated the deeper purification of cer-
tain recalcitrant organic compounds, and this process
would occur in the saturated aquifer layer. Therefore, an
adequate retention time is especially important for the
removal of refractory materials, which is consistent with
previous laboratory and field studies that monitored the
removal characteristics of organic matter (Grunheid et al.,
2005; Zhang et al., 2007).
Differences in microbial functional diversity among the
soil samples were compared using mean Shannon–Weaver
indices (Table 2). All three indices (diversity H′, richnessR and evenness J′) of sample S1 were significantly higher
than those of samples S2–S4, whereas values for S2–S4were similar. These results clearly indicate that the micro-
bial community associated with S1 had a higher func-
tional diversity than the other microbial communities.
The functional differences among the four communi-
ties were analysed in detail using PCA based on the
absorbance data for the 31 individual carbon substrates at
72.6 h (Fig. 5 and Table S1). Clear separation of S1 and
the other three soil samples (S2, S3 and S4) were detected
based on PC1, which explains 45.0% of the variance.
According to the study of Garland & Mills (1991), the
most important carbon sources in differentiating among
the communities were defined as those that had at least
half of their variance explained by PCs (Table 3). On the
basis of PC1, S1 had positive coordinates, whereas S2–S4had negative coordinates. The microbial community in S1
utilized several carbohydrates (D-xylose, D-cellobiose,
i-erythritol, methyl b-D-glucoside, a-D-lactose, D-mannitol
and N-acetyl-D-glucosamine), carboxylic acid (2-hydroxy-
benzoic acid), amino acids (L-serine), polymers (glycogen,
a-cyclodextrin) and amines (phenylethylamine) to a
greater degree, and several carboxylic acids (c-hydroxybu-tyric acid, a-ketobutyric acid, D-galactonic acid lactone
and itaconic acid), phosphorylated chemicals (DL-a-glyc-erol phosphate), amino acids (L-threonine, L-phenylala-
nine) and esters (pyruvic acid methyl ester) to a lesser
degree compared with microbial communities associated
with S2–S4. PC2 explained 19.5% of the variance, and
coordinate values increased in the saturated aquifer along
the flow path (S2–S4). Separation of these soils largely
resulted from the greater utilization of esters (pyruvic
acid methyl ester), carboxylic acids (D-glucosaminic acid),
phosphorylated chemicals (glucose 1-phosphate) and
amino acids (glycyl L-glutamic acid) and lesser utilization
of amino acids (L-asparagine), carboxylic acids
(4-hydroxy benzoic acid, and itaconic acid), polymers
(tween 40) and amines (putrescine). These results indicate
that the functional structures of the microbial community
remained steady in the saturated aquifer, consistent withFig. 5. Score plots of the PCA based on 72.6 h absorbance data.
Table 3. Correlation of carbon source variables to PCs
PC1 PC2
Carbon source r* Carbon source r*
Amines Amines
Phenylethylamine 0.92 Putrescine �0.54
Amino acids Amino acids
L-Serine 0.69 Glycyl L-glutamic
acid
0.87
L-Threonine �0.89 L-Asparagine �0.90
L-Phenylalanine �0.65 Carboxylic acids
Carbohydrates D-Glucosaminic acid 0.72
D-Cellobiose 0.92 4-Hydroxy benzoic
acid
�0.88
D-Xylose 0.90 Itaconic acid �0.61
i-Erythritol 0.85 Esters
Methyl b-D-Glucoside 0.73 Pyruvic acid methyl
ester
0.74
a-D-Lactose 0.67 Phosphorylated chemicals
D-Mannitol 0.58 Glucose
1-phosphate
0.74
N-Acetyl-D-glucosamine 0.50 Polymers
Carboxylic acid Tween 40 �0.75
2-Hydroxybenzoic acid 0.84
c-Hydroxybutyric acid �0.99
a-Ketobutyric acid �0.97
D-Galactonic acid
lactone
�0.79
Itaconic acid �0.69
Esters
Pyruvic acid methyl
ester
�0.63
Phosphorylated chemicals
DL-a-Glycerol phosphate �0.90
Polymers
Glycogen 0.86
a-Cyclodextrin 0.82
*Pearson’s regression coefficient.
FEMS Microbiol Ecol 80 (2012) 9–18 ª 2011 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
Microbial dynamic during SAT for water recharge 15
the result showing slight variation of bulk organic com-
pounds found in the recharged water of the saturated soil
aquifer.
EEM spectra of the water samples
The different compositions of DOMs in the water sam-
ples were identified in the EEM spectra and were quanti-
fied as the normalized region-specific excitation–emission
area volumes (ui,n) using FRI (Chen et al., 2003). The
fluorescence intensity in all five regions decreased signifi-
cantly after ozonation, with approximately 60% removal
of total ui,n. In particular, removal of fulvic acid-like
(Region III, Ex < 250 nm, Em > 380 nm) and humic
acid-like compounds (Region V, Ex > 280 nm, Em >380 nm) was relatively higher (approximately 70%). By
contrast, around 30% of total ui,n was removed during
SAT, with significantly higher removal of 60% on aro-
matic proteins (Region I, Ex < 250 nm, Em < 330 nm)
and 40% on soluble microbial byproduct-like compounds
(Region IV, Ex = 250–280 nm, Em < 380 nm). Clearly,
differences in the removal preferences of fluorescent
DOMs were determined between ozonation and SAT,
indicating complementary effects in this treatment system.
Correlation between metabolic activity and
nutrients
Microbial functional diversity (e.g. H′, R, J′) and biomass
(e.g. VS, phospholipids) of soil are considered to be
related to the water quality (e.g. DOC, UV254) of the
respective feed water. These correlations were investigated
using the Pearson product-moment correlation. DOC was
correlated significantly (P < 0.05) with average metabolic
rates, R, VS and PLE results. Other indicators of water
quality (i.e. UV254, N, P) showed no significant correla-
tion with microbial biomass or functional diversity
(P > 0.05). This result indicates that organic carbon con-
tent is the key limiting factor for development of micro-
organisms, which can be explained by C/N/P ratios.
Specifically, aliphatic compounds contributed more to
microbial community development than aromatic com-
pounds. Moreover, microbial biomass (VS and phospho-
lipids), diversity (R) and metabolic rate were significantly
correlated with each other, thus confirming the reliability
of the BIOLOG methods.
To gain deeper insight into the effects of DOM compo-
sitions on the microbial metabolic activity during SAT,
correlations between the metabolic rates of various carbon
substrates and Pi,n of various DOMs in the respective feed
water were analysed using the Pearson product-moment
correlation. Tyrosine-like aromatic proteins in Region I
and soluble microbial byproduct-like material in Region
IV were significantly correlated with the oxidation rates of
most carbon source groups (i.e. polymers, carbohydrates,
amino acids, amines and phosphorylated chemicals)
(P < 0.05). These findings agree with the results above,
showing that the preferential removal of Regions I and IV
was observed during SAT. However, the oxidation rates
of carboxylic acids and esters were not significantly corre-
lated with any of the fluorescent materials in the feed
water (P > 0.05), which may be attributable to the non-
fluorescent characteristics of responsible substrates. Fulvic
acid-like (Region III) and humic acid-like compounds
(Region V) were negatively correlated with all the oxida-
tion rates, which correspond to their refractory character-
istics in SAT (Drewes & Jekel, 1998; Maeng et al., 2011).
In conclusion, biomass analysis and BIOLOG assay,
coupled with ESEM, enable the dynamics of microbial
communities during SAT to be studied. Regarding the
fate of contaminants, soil biomass and functional diver-
sity in the unsaturated vadose layer were significantly
higher than those in the saturated aquifer, whereas they
remained at the same level along the saturated aquifer.
The vadose layer soil sample was demonstrated to be
clearly separated from the saturated layer samples using
PCA based on substrate utilization patterns. Exception-
ally, the oxidation rates of esters remained steady during
SAT, indicating the purification potential of the saturated
aquifer on certain recalcitrant organic compounds given
an adequate retention time.
Acknowledgements
This research was supported by Major Science and Tech-
nology Program for Water Pollution Control and Treat-
ment (Grant no. 2008ZX07314-008-04) and the National
Natural Science Foundation of China (Grant nos.
50878115 and 51078211).
References
Al-Mutairi NZ (2009) Variable distributional characteristics of
substrate utilization patterns in activated sludge plants in
Kuwait. Bioresour Technol 100: 1524–1532.APHA 2540 (2005) Standard methods for the examination of
water and wastewater. American Public Health Association/
American Water Work Association/Water Environment
Federation, 21st edn (Eaton AD, Clesceri LS & Greenberg
AE, eds), pp. 60–61. APHA, Washington, DC.
Chen W, Westerhoff P, Leenheer JA & Booksh K (2003)
Fluorescence excitation emission matrix regional integration
to quantify spectra for dissolved organic matter. Environ Sci
Technol 37: 5701–5710.Conn KE, Siegrist RL, Barber LB & Meyer MT (2010) Fate of
trace organic compounds during vadose zone soil treatment
ª 2011 Federation of European Microbiological Societies FEMS Microbiol Ecol 80 (2012) 9–18Published by Blackwell Publishing Ltd. All rights reserved
16 X. Zhang et al.
in an onsite wastewater system. Environ Toxicol Chem 29:
285–293.Drewes JE & Fox P (1999) Fate of natural organic matter
(NOM) during groundwater recharge using reclaimed water.
Water Sci Technol 40: 241–248.Drewes JE & Jekel M (1998) Behavior of DOC and AOX using
advanced treated wastewater for groundwater recharge.
Water Res 32: 3125–3133.Firestone M, Balser T & Herman D (1998) Defining Soil
Quality in Terms of Microbial Community Structure. Annual
Reports of Research Projects, University of California,
Berkeley, Berkeley, CA.
Fox P, Aboshanp W & Alsamadi B (2005) Analysis of soils to
demonstrate sustained organic carbon removal during soil
aquifer treatment. J Environ Qual 34: 156–163.Garland JL & Mills AL (1991) Classification and
characterization of heterotrophic microbial communities on
the basis of patterns of community-level sole carbon-source
utilization. Appl Environ Microbiol 57: 2351–2359.Gomez E, Ferreras L & Toresani S (2006) Soil bacterial
functional diversity as influenced by organic amendment
application. Bioresour Technol 97: 1484–1489.Grossart HP & Simon M (1998) Bacterial colonization and
microbial decomposition of limnetic organic aggregates
(lake snow). Aquat Microb Ecol 15: 127–140.Grunheid S, Amy G & Jekel M (2005) Removal of bulk
dissolved organic carbon (DOC) and trace organic
compounds by bank filtration and artificial recharge. Water
Res 39: 3219–3228.Harvey RW, Smith RL & George L (1984) Effect of organic
contamination upon microbial distributions and
heterotrophic uptake in a Cape Cod, Mass. Aquifer. Appl
Environ Microbiol 48: 1197–1202.Hendel B, Marxsen J, Fiebig D & Preuss G (2001) Extracellular
enzyme activities during slow sand filtration in a water
recharge plant. Water Res 35: 2484–2488.Jena S, Jeanmeure LFC, Wichukorn SD & Wright PC (2006)
Carbon substrate utilisation profile of a high concentration
effluent degrading microbial consortium. Environ Technol
27: 863–873.Joyce E, Phull SS, Lorimer JP & Mason TJ (2003) The
development and evaluation of ultrasound for the treatment
of bacterial suspensions. A study of frequency, power and
sonication time on cultured Bacillus species. Ultrason
Sonochem 10: 315–318.Kolehmainen RE, Langwaldt JH & Puhakka JA (2007)
Natural organic matter (NOM) removal and structural
changes in the bacterial community during artificial
groundwater recharge with humic lake water. Water Res
41: 2715–2725.Kolehmainen RE, Tiirola M & Puhakka JA (2008) Spatial and
temporal changes in Actinobacterial dominance in
experimental artificial groundwater recharge. Water Res 42:
4525–4537.Kolehmainen RE, Korpela JP, Munster U, Puhakka JA &
Tuovinen OH (2009) Extracellular enzyme activities and
nutrient availability during artificial groundwater recharge.
Water Res 43: 405–416.Liu WJ (2003) Study on Characteristics of Biodegradable
Organic Matter and Disinfection By-Products in Drinking
Water. Higher Education Press, Beijing.
Maeng SK, Sharma SK, Lekkerkerker-Teunissen K & Amy GL
(2011) Occurrence and fate of bulk organic matter and
pharmaceutically active compounds in managed aquifer
recharge: a review. Water Res 45: 3015–3033.Middelboe M, Søndergaard M, Letarte Y & Borch NH (1995)
Attached and free-living bacteria: production and polymer
hydrolysis during a diatom bloom. Microb Ecol 29:
231–248.Miller GW (2006) Integrated concepts in water reuse:
managing global water needs. Desalination 187: 65–75.Ministry of Environmental Protection PR China (2002)
Standard Methods of Water and Wastewater Monitoring of
China, 4th edn. China Environmental Science Press, Beijing.
Preston-Mafham J, Boddy L & Randerson PF (2002) Analysis
of microbial community functional diversity using sole-
carbon-source utilisation profiles – a critique. FEMS
Microbiol Ecol 42: 1–14.Quanrud DM, Hafer J, Karpiscak MM, Zhang HM, Lansey KE
& Arnold RG (2003) Fate of organics during soil-aquifer
treatment: sustainability of removals in the field. Water Res
37: 3401–3411.Rauch T & Drewes JE (2005) Quantifying biological organic
carbon removal in groundwater recharge systems. J Environ
Eng 131: 909–923.Rauch-Williams T & Drewes JE (2006) Using soil biomass as
an indicator for the biological removal of effluent-derived
organi carbon during soil infiltration. Water Res 40:
961–968.Schutz K, Kandeler E, Nagel P, Scheu S & Ruess L (2010)
Functional microbial community response to nutrient pulses
by artificial groundwater recharge practice in surface soils
and subsoils. FEMS Microbiol Ecol 72: 445–455.Vanderzalm JL, Le Gal La Salle C & Dillon PJ (2006) Fate of
organic matter during aquifer storage and recovery (ASR) of
reclaimed water in a carbonate aquifer. Appl Geochem 21:
1204–1215.Wang C, Xi JY, Hu HY & Yao Y (2009) Effects of UV
pretreatment on microbial community structure and
metabolic characteristics in a subsequent biofilter treating
gaseous chlorobenzene. Bioresour Technol 100: 5581–5587.Weber KP, Gehder M & Legge RL (2008) Assessment of
changes in the microbial community of constructed wetland
mesocosms in response to acid mine drainage exposure.
Water Res 42: 180–188.Xue S, Zhao QL, Wei LL & Ren NQ (2009) Behavior and
characteristics of dissolved organic matter during column
studies of soil aquifer treatment. Water Res 43: 499–507.Zhang ZY, Lei ZF, Zhang ZY, Sugiura N, Xu XT & Yin DD
(2007) Organics removal of combined wastewater through
shallow soil infiltration treatment: a field and laboratory
study. J Hazard Mater 149: 657–665.
FEMS Microbiol Ecol 80 (2012) 9–18 ª 2011 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved
Microbial dynamic during SAT for water recharge 17
Zhao X, Zhang M & Cheng XZ (2009) Bulk organic matter
and nitrogen removal from reclaimed water during
groundwater recharge by enhanced direct injection well.
Water Environ Res 81: 69–75.
Supporting Information
Additional Supporting Information may be found in the
online version of this article:
Table S1. Carbon sources that could be utilized by differ-
ent column soil samples.
Please note: Wiley-Blackwell is not responsible for the
content or functionality of any supporting materials sup-
plied by the authors. Any queries (other than missing
material) should be directed to the corresponding author
for the article.
ª 2011 Federation of European Microbiological Societies FEMS Microbiol Ecol 80 (2012) 9–18Published by Blackwell Publishing Ltd. All rights reserved
18 X. Zhang et al.