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RESEARCH ARTICLE
Biodegradation of a biochar-modified waterborne polyacrylate
membrane coating for controlled-release fertilizer
and its effects on soil bacterial community profiles
Zijun Zhou &Changwen Du &Ting Li &Yazhen Shen &
Yin Zeng &Jie Du &Jianmin Zhou
Received: 5 October 2014 / Accepted: 22 December 2014
#Springer-Verlag Berlin Heidelberg 2015
Abstract Biochar-modified polyacrylate-like polymers are
promising waterborne polymer-based membrane coatings for
controlled-release fertilizers. However, the effect of these
membrane polymers on paddy soil is unknown. A soil incu-
bation experiment was conducted using Fourier transform in-
frared photoacoustic spectroscopy to monitor the changes in
the polymer-coated membranes in paddy soil, and Biolog
EcoPlates and polymerase chain reaction-denaturing gradient
gel electrophoresis were used to detect the effects of the mem-
branes on soil bacterial community profiles. Compared to un-
modified membranes, the biodegradation rate of the biochar-
modified membrane was slower, and the membrane was more
intact, which improved and guaranteed the controlled release
of nutrients. Compared to the soil without membranes, the
biochar-modified membranes, as well as unmodified ones,
showed no significant impacts on the composition diversity
of soil dominant bacterial community. The activity and func-
tional diversity of soil culturable microbial community during
the early stage of incubation were reduced by biochar-
modified membranes due to the release of small amount of
soluble organic materials but were both recovered in the 12
th
month of the incubation period. Therefore, the biochar-
modified waterborne polyacrylate was environmentally
friendly, demonstrating its potential both in the development
of coated controlled-release fertilizers and in the utilization of
crop residue.
Keywords Biochar .Waterborne polyacrylate .
Biodegradation .Bacteria .Controlled-release fertilizer
Introduction
Fertilizers are extremely important for crop yield, and
controlled-release fertilizers (CRFs) coated with a polymer
control the release of nutrients, so their availability coincides
with the crop’s requirement, which shows great potential in
agriculture (Shaviv 2001). In contrast to conventional fertil-
izers, CRFs are advantageous because they reduce nutrient
leaching by rain or irrigation and save labor-associated costs
of fertilizer application (Han et al. 2009;Zhaoetal.2010),
especiallyfor rice production (Choudhury and Kennedy 2005;
Li et al. 2009;Yangetal.2012).
Control over the rate of nutrient release from coated fertil-
izer is mainly decided by the polymer coating. Currently, there
are two main kinds of coating polymers, and they are distin-
guished by the type of solvent used in CRF production. There
are the organic-solvent-dissolvable polymers, such as poly-
acrylamide (Rajsekharan and Pillai 1996) or polystyrene
(Garcia et al. 1996), and most of them are expensive and toxic
to the environment in production and in application. The other
type of coating polymer is the water-dissolvable polymer,
such as the waterborne polymers, which are relatively cheap
and environmentally nontoxic during production (Zhao et al.
2010). Consequently, increasing interests are focused on wa-
terborne polymer coatings.
Responsible editor: Robert Duran
Electronic supplementary material The online version of this article
(doi:10.1007/s11356-014-4040-z) contains supplementary material,
which is available to authorized users.
Z. Zhou :C. Du (*):T. Li :Y. Sh en :Y. Zeng :J. Du :J. Zhou
State Key Laboratory ofSoil and Sustainable Agriculture, Institute of
Soil Science, Chinese Academy of Sciences, 210008 Nanjing, China
e-mail: chwdu@issas.ac.cn
Z. Zhou :T. L i :Y. Z e n g :J. Du
University of Chinese Academy of Sciences, 100049 Beijing, China
T. Li
College of Resources and Environment, Sichuan Agricultural
University, 611130 Chengdu, China
Environ Sci Pollut Res
DOI 10.1007/s11356-014-4040-z
Degradation of the CRF polymer coating in soil is a factor
that is highly considered for control over the rate of nutrient
release and also for environmental safety. Degradation of the
polymerisinfluencedbysoiltemperature,bysoilmoisture,
and especially by soil biological activity (Kennedy and Smith
1995;Shahetal.2008). Degradation of the polymer refers not
only to bond scission and chemical transformation but also to
the formation of new functional groups (Celina 2013). Water-
borne polyacrylate contains ester groups and carboxyl groups
which are sensitive to moisture (Zhang et al. 2012). These
groups offer opportunities to tune the biomechanical proper-
ties within a broad range of desired properties and to acceler-
ate polymer degradation (Decker and Zahouily 1999;Zhang
et al. 2012). Variation in polymer degradation is increased by
the complex soil environment, which may result in polymer
coating failure and unguaranteed control over the rate of nu-
trient release. On the other hand, the degradation products of
synthetic polymers may impact on microorganisms that play a
central role in nutrient cycling and provide an important eco-
system service (Albertsson and Karlsson 1990;Costanzaetal.
1987; Hadad et al. 2005). Furthermore, changes in soil bacte-
rial community profiles are important to the soil environment
(Kirk et al. 2004). For example, Liu et al. (2011)detectedthat
one kind of CRF resin coating increased the quantity of
bacteria and actinomyces, determined using plate count and
enzyme activity in soil, and Ikeda et al. (2014) found that urea-
formaldehyde, the oldest slow-release N fertilizer, markedly
increased bacterial diversity. Despite waterborne polymer-
coated CRFs having the most potential, there is little published
information about changes to soil bacterial profiles in re-
sponse to their degradation in soil.
In our previous study, we found that waterborne
polyacrylate emulsion could be applied to CRF development;
furthermore, biochar-modified waterborne polyacrylate mate-
rial enhanced the mechanical strength and increased the re-
lease period, hence demonstrating great potential for water-
borne polymer-coated CRFs (Zhou et al. 2013). The biochar
involved was derived from locally available wheat residues
(Huggins et al. 2014;Lehmann2007), and the effect of the
biochar on soil microbial profiles was studied (Zimmerman
et al. 2011). However, the degradation of biochar-modified
waterborne polyacrylate membranes and its impact on soil
microbes are unknown.
The objectives of this study were (1) to detect the degrada-
tion of the biochar-modified waterborne polyacrylate mem-
brane in paddy soil using Fourier transform infrared photo-
acoustic spectroscopy (FTIR-PAS) and (2) to explore the ef-
fects of biochar-modified waterborne membrane on com-
position diversity of the soil dominant bacterial commu-
nity and functional diversity of the soil culturable bacte-
rial community by polymerase chain reaction-denaturing
gradient gel electrophoresis (PCR-DGGE) and Biolog
EcoPlates, respectively.
Materials and methods
Materials
Waterborne polyacrylate emulsion (Doctor Hydrophilic
Chemicals Co., Ltd., Yizheng, China) was used as a coating
material to control the rate of nutrient release. The emulsion
(50 % dry matter content) contained butyl acrylate, methyl
methacrylate, methyl acrylic acid, and the cross-linker
aziridine. The biochar was from wheat straw that was pyro-
lyzed at 400 °C; detailed information has been presented else-
where (Xu et al. 2013). Biochar (1 % w/w) was added to the
waterborne polyacrylate emulsion for both chemical and
physical modification, i.e., 0.275-g crushed biochar was
added into 55-g raw emulsion and then stirred for 15 min at
room temperature. The detailed information about making
membranes was as follows: 55-g waterborne polyacrylate la-
tex or biochar-modified waterborne polyacrylate latex was
distributed into a polytef culture dish (internal diameter,
10 cm) dried in an oven at 80 °C for 2 h and formed into the
membranes of 1-mm thickness on dish surface; then, the mod-
el membranes removed from the mold were tailored into cir-
cles with 1-cm diameter and stored in a 4 °C refrigerator for
use.
Paddy soil was collected from the Ecological Station of
Red Soil, Chinese Academy of Sciences, in Yingtan City,
Jiangxi Province of China (28° 15′N, 116° 55′E). Soil
agro-chemical properties were as follows: pH (H
2
O), 5.2; or-
ganic carbon, 22.13 g kg
−1
; total nitrogen, 1.92 g kg
−1
;total
phosphorous, 0.61 g kg
−1
;andtotalpotassium,6.01gkg
−1
.
The soil texture used in the experiment was loam soil (Ultisols
and Oxisols in US Soil Taxonomy) that contained 38 % sand,
42 % silt, and 20 % clay. The soil was air-dried at room
temperature and passed through a 2-mm sieve.
Soil incubation experiment
Twenty-seven of polyethylene cups (height, 13 cm; internal
diameter, 8.5 cm) were filled with 500 g of soil, and four
pieces of membrane (about 0.7 g) were buried horizontally
in the soil at 1-cm depth from the soil surface. The soil was
waterlogged, and a 2-cm layer of water was maintained on the
surface. Deionized water was used for all treatments. The cups
were covered with plastic film perforated with holes for gas
exchange and incubated at 28±1 °C. Water was added to the
cups every 2 days to maintain the water layer.
Bifactorial design with the factors treatment and incubation
time each replicated for three times was set. The first factor
was membrane treatment which included control without
membranes in soil (CK), unmodified waterborne polyacrylate
(UP) membranes, and biochar-modified waterborne
polyacrylate (BP) membranes. The second factor was sam-
pling position treatment which included the contacted soil
Environ Sci Pollut Res
layer (top layer, 0–2-cm distance from the buried membrane
surface) and the noncontacted soil layers (bottom layer, 8–10-
cm distance from the buried membrane surface). Soil samples
and buried membranes were both sampled after 2, 6, and
12 months.
The sampled membranes were washed with water and
dried at 80 °C, weighed, and then measured using FTIR-
PAS. The sampled fresh soil was divided into three subsam-
ples. One was stored at 4 °C for detecting the soil physical and
chemical properties; the second was used to determine the
functional diversity of soil culturable microbial communities
by Biolog EcoPlates; the third was stored at −20 °C for the
evaluation of the genetic structures of soil dominant bacteria
using PCR-DGGE.
Recording of FTIR-PAS spectra
FTIR-PAS was based on the photoacoustic effect while the
photoacoustic signal generation was affected by the physical
properties of soils (Rosencwaig and Gersho 1976), and the
technique has been previously demonstrated to be very suit-
able to analyze polymeric materials, especially the heteroge-
neous compound polymers without sample pretreatments
when compared with conventional transmission and reflection
techniques (Almeida et al. 2002;Duetal.2010; Zhang et al.
2012). FTIRspectroscopyhas been used widely to qualify and
quantify the polymer degradation (Merlatti et al. 2008;Perrin
et al. 2009; Zhang et al. 2013). FTIR-PAS spectra of the mem-
branes were recorded using an FTIR spectrometer (Nicolet
6700) with a photoacoustic accessory (MTEC model 300,
USA). The scans were conducted in the wavenumber range
of 500–4000 cm
−1
with resolution of 4 cm
−1
, using 32 scans
and mirror velocities of 0.16, 0.32, 0.64, and 1.28 cm s
−1
.For
the spectra recording, a piece of the membrane was put into
the photoacoustic accessory cell, and the cell was purged with
dry helium for 10 s prior to scanning. Black carbon was used
as reference (Du et al. 2010).
The profiling depth of the membrane was obtained using
FTIR-PAS according to Eq. (1):
μ¼ffiffiffiffiffiffiffiffi
D
πvγ
sð1Þ
where μis the profiling depth (μm), Dis the thermal diffusiv-
ity of sample, vis the moving mirror velocity (cm s
−1
), and γis
the wavenumber (cm
−1
). The thermal diffusivity of most poly-
meric materials was about 0.01×10
−5
m
2
s
−1
(Zhang et al.
2012), and the profiling depths were calculated under different
moving mirror velocities and wavenumbers (Table 1).
Principal component analysis was conducted on the FTIR-
PAS spectra, and the Euclidean distances using the first 11
components were used to determine dissimilarities between
membranes before incubation and membranes in each
incubation period. Different spectral ranges, i.e., 500–4000,
2800–3200, 1500–1900, 1300–1500, 1000–1300, and 500–
1000 cm
−1
, were selected according to functional group to
calculate the Euclidean distance. A higher Euclidean distance
meant greater compositional difference between membranes.
Determination of soil physical and chemical properties
Soil pH was measured after shaking a soil-water (1:5w/v)
suspension for 30 min. Soil organic carbon (SOC) was deter-
mined by dichromate oxidation and titration with ferrous am-
monium sulfate (Walkley and Black 1934). A soil-water (1:5
w/v) suspension was shaken at 200 rev min
−1
for 1 h, then
centrifuged at 4000 rpm for 20 min, and the centrifuged su-
pernatant which was filtered through sterile 0.45-μmsyringe
filters was used to detect the soil dissolved organic carbon
(DOC) and dissolved organic nitrogen (DON) on a total or-
ganic carbon analyzer (Multi N/C 3000, Germany).
Biolog EcoPlate analysis
Biolog EcoPlates (Biolog, Inc., USA) have been widely used
to determine the total activity and functional diversity of soil
culturable microbial communities (Harch et al. 1997). The 96-
well EcoPlates contained 31 different carbon sources, replicat-
ed for three times, plus three blank wells without any carbon
source. A 5-g sample of fresh soil was suspended in 50 mL of
0.85 % (w/v) sterile NaCl solution, shaken for 30 min, and
then allowed to settle for 5 min. The supernatant was diluted
10-fold. A 150-μL aliquot of the diluted sample was inoculat-
ed onto the EcoPlate and incubated at 28 °C in the dark.
Substrate utilization on the plate was monitored by measuring
the absorbance at 590-nm wavelength every 24 h for 168 h
using a Bio-Rad Microplate Reader (Bio-Rad, USA). The
total culturable microbial activity and functional diversity
were measured by the average well color development
(AWCD) and three diversity indices (Shannon-Weaver index
H′, Simpson index 1/D, and McIntosh index U)accordingto
Fang et al. (2012).
PCR-DGGE analysis
To assess changes in composition diversity of the soil domi-
nant bacterial community, a PCR-DGGE method was used
(Muyzer et al. 1993). Soil DNA was extracted from each ho-
mogenized sample (approximately 0.5 g) following the man-
ufacturer’s protocols using the FastDNA® SPIN Kit for soil
(MP, USA). The extracted soil genomic DNAwas dissolved in
50-μL TE buffer and stored at −20 °C prior to use. The DNA
samples were purified using PowerClean® DNA Clean-Up
Kit (MO BIO, Inc., USA), and PCRs were run using 50-μL
reaction volumes. The PCR procedures were as follows: an
initial94°Cdenaturationfor5min,followedby35cyclesat
Environ Sci Pollut Res
94 °C for 30 s, 60 °C for 30 s, 72 °C for 30 s, and a final
extension step at 72 °C for 7 min, and then held at 4 °C. Using
these conditions, the PRBA338F (5′-ACT CCT ACGGGA
GGC AGC AG-3′) and PRUN518R (5′-ATT ACC GCG
GCT GCT GG-3′) primers were used to amplify the 338 to
518 ribosomal DNA (rDNA) region, as described by Nakatsu
et al. (2000), and to obtain products of about 200 bp, while
forward primers contained a 40-bp GC-clamp (5′-CGC CCG
CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG
GGG G-3′) attached to the 5′end (Muyzer et al. 1993).
Amplicons were checked by electrophoresis on 1 % agarose
gel. We then performed 16S rDNA-DGGE using the DCode
System (Bio-Rad, USA). Aliquot (10 μL) of amplicons was
loaded (top filling method) onto 8 % acrylamide-
bisacrylamide gel containing a denaturant gradient of 45 to
65 % at 70 V for 16 h in 1× TAE running buffer at 60 °C,
followed by 0.5-h coloration using Gel-Red nucleic acid gel
stain (Biotium, USA). The gels were visualized and digita-
lized by using a Gel Doc™EQ imager (Bio-Rad, USA) com-
bined with Quantity One 4.4.0 (Bio-Rad, USA). Species rich-
ness was calculated from the band numbers per sample. H′and
evenness were calculated using the number of bands and peak
intensities (Yu and Morrison 2004).
Statistical analysis
All spectral data were processed using Matlab 2009b. Digital
information for DGGE was determined using Quantity One
4.4.0. All bacterial community diversity data were analyzed
by a three-way analysis of variance (ANVOA) using SPSS
16.0 to reveal the main effects and potential interactions.
The assumption of homogeneity of variance was tested with
Levene’stestata=0.05. When statistically significant differ-
ences existed according to ANOVA (P<0.05), treatment
means were compared using Tukey test at a=0.05. Euclidean
distance changes of the two membranes over the incubation
period were detected by the Tukey test at the 0.05 probability.
A redundancy analysis (RDA) was used to detect the effects of
main soil properties on soil bacterial community in the vegan
package of R v.2.8.1 project (R Development Core Team.
2006). The graphics were plotted using SigmaPlot 12.5.
Results and discussion
The change of waterborne membranes overthe soil incubation
period
Figure 1a shows the weight loss of waterborne membranes
buried in the waterlogged paddy soil during the 12-month
incubation. Compared with the control, the weight losses for
the UP and BP membranes after 2 months were about 1.01
and 0.99 %, respectively, and increased to 1.24 and 1.15 %
after 12 months, respectively. This demonstrates that the
membranes were degraded at a relatively low rate; thus, they
remained integrated and could function effectively as
Tabl e 1 The profiling depth of polymer membrane using FTIR-PAS calculated using typical thermal diffusivity, wavenumber, and moving mirror
velocity
Assignment Band position (cm
−1
)Moving-mirror
velocity (cm s
−1
)
Thermal diffusion
distance (μm)
C–H stretching vibration ~2900 0.16 8.21
0.32 5.81
0.64 4.11
1.28 2.90
C═O stretching vibration ~1730 0.16 10.72
0.32 7.58
0.64 5.36
1.28 3.79
C–H bending vibration ~1450 0.16 11.72
0.32 8.28
0.64 5.86
1.28 4.14
C–O stretching vibration ~1100 0.16 13.10
0.32 9.26
0.64 6.55
1.28 4.63
The profiling depth function of photoacoustic spectroscopy is μ¼ffiffiffiffiffiffi
D
πνγ
q,inwhichμis the profiling depth (μm), Dis the thermal diffusivity of sample
(0.01×10
−5
m
2
s
−1
), vis the moving-mirror velocity (cm s
−1
), and γis the wavenumber (cm
−1
)
Environ Sci Pollut Res
controlled-release membranes. Figure 1b further demonstrates
that the UP and BP membranes remained totally intact. The
color of the UP membranes became darker over the incubation
period, suggesting that some small molecules in the soil might
diffuse into the membranes and that the membranes were
compatible with soil. The color of the BP membranes
remained black due to the biochar component, and only
a small amount of biochar was lost from the membrane
border according to the membrane color, suggesting that
BP was a more stable coating than UP for controlled
nutrient release.
Figure 2shows the spectra of all samples over the incuba-
tion period, and several similar function groups are apparent,
such as 2900, 1730, 1450, and 1200 cm
−1
which represent the
C–H stretching vibration, the C═O stretching vibration, the
C–Hbendvibration,andtheC–O stretching vibration, respec-
tively. However, there were numerous variances in the spectra
between treatments.
AccordingtoEq.(1), the profiling depths (μm) were
calculated at different moving mirror velocities (Table 1).
Figure 2shows the FTIR-PAS spectra of UP and BP mem-
branes at different moving mirror velocities and indicates
6 months 12 months
BP
UP
Incubation period (month)
0month 2months
2612
Weight loss (%)
0.8
1.0
1.2
1.4
UP
BP
ab
Fig. 1 Change in waterborne
membranes over 12 months of
incubation in paddy soil at 28 °C.
aWeight loss of membrane and b
visual surface morphology of
membrane. UP unmodified
polyacrylate membrane, BP
biochar-modified polyacrylate
membrane
100015002000250030003500
-1
0
1
2
3
4
0.16 cm-1 s-1
0.32 cm-1 s-1
0.64 cm-1 s-1
1.28 cm-1 s-1
100015002000250030003500
-1
0
1
2
3
4
0.16 cm-1 s-1
0.32 cm-1 s-1
0.64 cm-1 s-1
1.28 cm-1 s-1
100015002000250030003500
-1
0
1
2
3
4
0.16 cm-1 s-1
0.32 cm-1 s-1
0.64 cm-1 s-1
1.28 cm-1 s-1
100015002000250030003500
-1
0
1
2
3
4
0.16 cm-1 s-1
0.32 cm-1 s-1
0.64 cm-1 s-1
1.28 cm-1 s-1
a
b
c
d
Wavenumber (cm-1)
Photoacoustic unit
Fig. 2 FTIR-PAS depth profiling spectra of waterborne membranes with
the moving mirror velocities of 0.16, 0.32, 0.64, and 1.28 cm s
−1
.aRaw
unmodified polyacrylate membrane (UP), bunmodified polyacrylate
membrane (UP) after 12 months of incubation in paddy soil at 28 °C, c
raw biochar-modified polyacrylate membrane (BP), and dbiochar-
modified polyacrylate membrane (BP) after 12 months of incubation in
paddy soil at 28 °C. The arrow shows the significant change around
1050 cm
−1
after 12 months of incubation in paddy soil
Environ Sci Pollut Res
that both UP and BP membranes were heterogeneous. For
the UP membranes (Fig. 2a), the surface layer (2.92–
4.65 μm, with a moving velocity of 1.28 cm s
−1
)showed
the greatest difference to the next three deeper layers
(4.14–13.10 μm, with a moving velocity of 0.16, 0.32,
and0.64cms
−1
, respectively). The most significant differ-
ence occurred for absorption around 1030 cm
−1
,whichis
assigned to CO–O–CorC–O–C vibration (Movasaghi
et al. 2008), indicating that more bonds of CO–O–Cor
C–O–C were observed in the surface layer (0–4.65 μm).
After 12 months of incubation, the UP membranes were
still heterogeneous, and the difference mainly occurred in
the surface layer, and the vibration intensity of CO–O–Cor
C–O–C became significantly weaker (Fig. 2b), suggesting
that some of these bonds were broken. Although similar
results were observed for the BP membranes (Fig. 2c), the
vibration intensity of CO–O–CorC–O–Cshowedanop-
posite trend compared with that for the UP membranes.
The vibration of CO–O–CorC–O–C in the surface layer
of the BP membranes was prevented by the involvement of
biochar, and after 12 months of incubation, the prevention
might be removed due to the loss of biochar from the sur-
face layer, as indicated by the ~50-cm
−1
shift of the absorp-
tion band toward the direction of lower wavenumbers:
980 cm
−1
for BP membranes versus 1030 cm
−1
for UP
membranes (Fig. 2d). Therefore, less degradation of CO–
O–CorC–O–C bonds occurred for the BP membranes
over 12 months of incubation, which demonstrated that
they were more stable than the UP membranes.
FTIR-PAS-spectra-based Euclidean distance was used to
judge the changes in the membranes during different incuba-
tion times. Six regions (500–4000, 500–1000, 1000–1300,
1300–1500, 1500–1900, and 2800–3200 cm
−1
)wereselected
to calculate the Euclidean distances. Because the main chang-
es occurred in the surface layer (the spectra with moving mir-
ror velocity of 1.28 cm s
−1
), the spectra at this depth were used
to calculate the Euclidean distances (Table 2). Significant dif-
ferences were observed for the UP membranes in the total
500–4000-cm
−1
region over 12 months of incubation. The
main contribution resulted from the fingerprint region of
500–1000 cm
−1
, followed by two other regions of 1300–
1500 and 1500–1900 cm
−1
, and the remaining regions showed
less contribution. However, for the BP membranes, although
there was some change in the 500–1000-cm
−1
region, no sig-
nificant difference was found in the total 500–4000-cm
−1
re-
gion, which further verified that the BP membranes were more
stable than the UP membranes.
Combining the results of the membrane weight loss, mor-
phology, and FTIR analysis, biodegradation of the waterborne
membranes mainly occurred in the surface layer, and the bio-
degradation rate of the UP membranes was significantly great-
er than that of the BP membranes. The surface layer was
directly subjected to the environment and was thus more
easily degraded, especially the C–O–Cgroupsthatwereeasily
broken. The involvement of biochar might form a thin coating
outside the C–O–C groups, which could protect the bonds
from degradation, although the groups were released after
12 months of incubation due to removal of biochar from the
surface layer. In addition, it is possible that the biochar prod-
ucts contained some toxic substance that suppressed microbial
activity, which reduced the polyacrylate biodegradability by
soil microorganism (Zimmerman et al. 2011).
Effects of waterborne polyacrylate membranes on soil
bacterial community profiles
The absorbance values at 96-h incubation from the Biolog
EcoPlates were used to evaluate the soil culturable bacterial
community functional diversity based on the AWCD value
and three diversity indices (Shannon-Weaver index H′,
Simpson index 1/D, and McIntosh index U). The AWCD,
which reflects the oxidative capacity on 31 kinds of carbon
sources in Biolog EcoPlates, is used as an indicator of overall
culturable microbial activity (Bossio and Scow 1995;Garland
1996). Three diversity indices (Shannon-Weaver index H′,
Simpson index 1/D, and McIntosh index U) were used to
assess the richness, dominant population, and evenness of soil
microorganisms, respectively (Fang et al. 2012). The AWCD,
H′,1/D,andUvalues of all samples at 96-h incubation were
significantly affected by membrane materials and incubation
period, and significant interaction variedly occurred among
Tabl e 2 Euclidean distances calculated from FTIR-PAS spectra data
over the incubation time at the surface layer with the moving mirror
velocity of 1.28 cm s
−1
Middle-infrared regions (cm
−1
) Materials Incubation period (month)
2612
500–4000
(whole spectral region)
UP 15.08c 21.51b 37.98a
BP 22.20a 17.80a 18.45a
2800–3200
(C–H stretching vibration)
UP 5.39a 6.53a 5.95a
BP 5.33a 5.17a 5.55a
1500–1900
(C═O stretching vibration)
UP 3.70b 5.23a 3.13b
BP 4.85b 5.73ab 7.41a
1300–1500
(C–H bending vibration)
UP 3.38b 5.6 a 3.42b
BP 3.19a 4.24a 3.59a
1000–1300
(C–O stretching vibration)
UP 3.13a 4.14a 2.79a
BP 3.04a 4.4 a 4.63a
500–1000
(fingerprint region)
UP 6.05b 9.68b 17.17a
BP 10.53a 8.82a 10.32a
Means within a row followed by the same letter are not significantly
different (P<0.05; Tukey test)
UP unmodified polyacrylate membrane, BP biochar-modified
polyacrylate membrane
Environ Sci Pollut Res
three factors (Table S1). The four functional diversity values
were sharply reduced at early 6 months of incubation period
and then gradually decreased (Tables 3and S1). It is possible
that aerobic microorganisms predominated in the early incu-
bation phase and then were suppressed, while anaerobic mi-
croorganisms predominated during the later incubation period
as oxygen was rapidly depleted (Liesack et al. 2000). In addi-
tion, products of anaerobic metabolism in soil, such as H
2
S,
NH
3
, or volatile fatty acids, may have inhibited microbial
activity (Sahrawat 2004), and the soil microorganisms gradu-
ally adapted to the conditions during the later phase.
In the second month, the AWCD values for both the top
and bottom layer in the CK and UP treatments were not sig-
nificantly different, but they were significantly higher than the
BP treatment, and the AWCD value for the top layer was
higher than that for the bottom layer for the BP treatment,
which means that the BP membrane suppressed the culturable
microbial activity both in the top layer and in the bottom layer
in the second month, and the suppression in the bottom layer
was alleviated. In the sixth month, the soil culturable micro-
bial activity was still suppressed by the BP treatment com-
pared with in top layer of the CK treatment, but the AWCD
value of the BP treatment was relatively less suppressed than
that in the second month, and the AWCD values in the bottom
layer for all treatments showed no significant difference, indi-
cating that the soil microbial activity in the bottom layer of the
BP treatment could be recovered. In the twelfth month, the
AWCD values in the top and bottom layers for all treatments
were not significantly different, indicating that the suppression
of soil microbial activity by biochar disappeared.
Tab le 3shows the changes of three diversity indices be-
tween treatments at each incubation period. For the top layer,
each of the three functional diversity indices of soil microbial
communities in the BP treatment was significantly lower in
the second month, but the McIntosh index Urecovered to the
CK level in the sixth month, and the Simpson index 1/Dand
Shannon-Weaver index H′recovered in the twelfth month,
indicating that richness, dominant population, and evenness
of soil microorganisms in the second month decreased in the
BP treatment, but the suppression of soil microorganism even-
ness disappeared in the sixth month, and the suppression of
dominant population and richness disappeared in the twelfth
month. For the bottom layer, the McIntosh index Uwas sig-
nificantly lower in the BP treatment, while the other two indi-
ces, i.e., Shannon-Weaver index H′and Simpson index 1/D,
showed no significant difference from the CK treatment in all
bottom layers in the second month, suggesting that the even-
ness of soil microorganisms was affected, but no differences
were observed for the index of the richness and dominant
population. Furthermore, there were no significant differences
within these three indices between the UP and CK treatments
over the incubation period.
According to previous studies (Peterson et al. 2013; Shen
et al. 2012;Wangetal.2011) and the variety of soil properties
among treatments in this study, the parameters of SOC, pH,
DOC, and DON were selected to do RDA analysis, which was
used to evaluate their effects on soil culturable bacterial activ-
ity and functional diversity. Figure 3shows that soil DOC and
DON have stronger effects (longer arrow) on the activity and
functional diversity of soil culturable bacterial community.
Therefore, two mechanisms may result in lower microbial
carbon utilization in the BP treatment during the early phase.
First, the biochar products contained some substances, such as
dioxins, furans, phenols, and polyaromatic hydrocarbons,
which could reduce the microbial activity (Zimmerman et al.
2011); second, a small amount of soluble fraction in the
Tabl e 3 Functional diversity indices of culturable microbial communities in paddy soil using Biolog EcoPlate over the soil incubation
Ave r ag e w ell c olor
development (AWCD)
Shannon-Wiener index H′Simpson index 1/DMcIntosh index U
Materials Sampling position Incubation period
(month)
Incubation period
(month)
Incubation period
(month)
Incubation period
(month)
2 6 12 2 6 12 2 6 12 2 6 12
CK Top layer 1.00a 0.54a 0.30a 3.19a 2.86a 2.57a 23.04a 13.61a 11.46a 6.49a 4.14ab 2.74a
Bottom layer 0.95a 0.30b 0.28a 3.15a 2.58bc 2.40a 21.91a 11.29ab 9.95a 6.30a 2.54c 2.74a
UP Top layer 0.91a 0.54a 0.29a 3.18a 2.78ab 2.53a 22.38a 14.72a 11.31a 5.95a 4.33a 2.62a
Bottom layer 0.95a 0.41ab 0.23a 3.21a 2.56bc 2.51a 23.65a 11.53ab 10.89a 6.07a 3.75ab 2.20a
BP Top layer 0.51c 0.35b 0.27a 2.89b 2.43c 2.37a 16.11b 9.79b 9.75a 3.92c 3.48abc 2.65a
Bottom layer 0.74b 0.35b 0.23a 3.12a 2.63bc 2.47a 20.90a 11.79ab 10.48a 5.01b 3.17bc 2.23a
Means within a column followed by the same letter are not significantly different (P<0.05; Tukey test)
CK soil without polyacrylate membrane, UP soil with unmodified waterborne polyacrylate membrane, BP soil with biochar-modified polyacrylate
membrane, top layer the contacted soil layer (soil with 0–2-cm distance from the buried membrane surface), bottom layer the noncontact soil layer (soil
with 8–10-cm distance from the buried membrane surface)
Environ Sci Pollut Res
membrane dissolved into the soil (Fig. 1), such as ammonium
persulfate catalyst, which lowered the carbon source utiliza-
tion (Li et al. 2013), and some organic materials of low mo-
lecular weight, which might affect the soil bacterial activity by
soil DOC and DON changes (Kiikkila et al. 2014). The sup-
pression of culturable microbial activity recovered over the
12-month incubation period, suggesting that short duration
of suppression due to the waterborne membrane involvement
might result in the proliferation of soil microorganisms.
The effects of waterborne polyacrylate membranes on
composition diversity of the soil dominant bacterial commu-
nity were verified by molecular analysis, which was per-
formed on DGGE gel using PCR amplification of 16S rDNA
genes from soil DNA of each sample and separated by
electrophoresis. Figure 4shows the soil dominant bacterial
community composition results on the DGGE gels in the
2
nd
,6
th
,and12
th
months. All samples showed that numerous
bacterial groups appeared to be ubiquitous (strong and weak),
thus indicating a polymicrobial community. The intensities
and numbers of bands for all samples reduced in the DGGE
gel, and the bands of DGGE lower gel became clear over the
incubation period. These results may be attributed to aerobic
microorganisms predominating during the early incubation
time, and they were then suppressed, while anaerobic micro-
organisms predominated over the incubation period as oxygen
was rapidly depleted (Liesack et al. 2000). The DGGE bands
were digitalized using Quantity One software for extracting
more information. Band numbers as well as intensities and
RDA1 (27.6 %)
RDA2 (6.5 %)
-0.8
-0.4
0.0
0.4
CK1
CK2
UP1
UP2
BP1
BP2
RDA1 (11.6 %)
-0.4 0.0 0.4 0.8 -0.4 0.0 0.4 0.8
RDA2 (5.7 %)
-0.8
-0.4
0.0
0.4
0.8
CK1
CK2
UP1
UP2
BP1
BP2
DOC
DON
pH
SOC
ba
pH
DOC
DONSOC
Fig. 3 Redundancy analysis (RDA) of the functional diversity of soil
culturable bacterial community with symbols coded by membrane
treatments. SOC soil organic carbon, DOC dissolved organic carbon,
DON dissolved organic nitrogen. aThe 2-month incubation and bthe
6-month incubation. CK soil without waterborne polyacrylate membrane,
UP soil with unmodified membrane, BP soil with biochar-modified
membrane, CK1,UP1,BP1 the contacted soil layer (soil with 0–2-cm
distance from the buried membrane surface) from treatment of CK, UP,
and BP, respectively, CK2,UP2,BP2 the noncontact soil layer (soil with
8–10-cm distance from the buried membrane surface) from treatment of
CK, UP, and BP, respectively
cba
CK1 CK2 UP1 UP2 BP1 BP2 CK1 CK2 UP1 UP2 BP1 BP2 CK1 CK2 UP1 UP2 BP1 BP2
Fig. 4 DGGE gel of bacterial community in paddy soil from different
treatments over the soil incubation. aThe 2-month incubation, bthe 6-
month incubation, and cthe 12-month incubation. CK soil without
waterborne polyacrylate membrane, UP soil with unmodified
membrane, BP soil with biochar-modified membrane, CK1,UP1,BP1
the contacted soil layer (soil with 0–2-cm distance from the buried
membrane surface) from treatment of CK, UP, and BP, respectively,
CK2,UP2,BP2 the noncontact soil layer (soil with 8–10-cm distance
from the buried membrane surface) from treatment of CK, UP, and BP,
respectively
Environ Sci Pollut Res
patterns provided information about the richness (S),
Shannon-Wiener diversity index (H′), and evenness (E),
which were calculated as follows: Operational taxonomic unit
(OTU) richness (S) was determined from the number of bands
in each lane, and the Shannon-Wiener diversity index (H′)was
calculated from H′=−∑P
i
lnP
i
, and evenness (E) was calculat-
ed as E=H′/H′
max
,whereH′
max
=lnS(Yu and Morrison 2004).
The indices for each treatment are listed in Table 4.According
to the three-way ANVOA, the most significant factor affecting
the indices was the incubation period (Table S2). During the
three sampling times, the OTU richness (S)values,Shannon-
Wiener index (H′), and evenness (E) values generally reduced
as a function of incubation time for all treatments (Tables 4
and S2). The anaerobic environment suppressed aerobic mi-
croorganisms (Liesack et al. 2000) and might contribute to the
reduced dominant bacterial community diversity in the three
treatments over the incubation period. And, in each sampling
time (2
nd
month, 6
th
month, and 12
th
month), three community
composition diversity values, i.e., S,H′,andEvalues, of BP
and UP treatments showed no significant difference from the
CK for both the top and bottom layers.
The results obtained by Biolog EcoPlates and PCR-
CGGE analyses are not necessarily in contrast, which is
similar to the results of others (Abbate et al. 2013;
Bushaw-Newton et al. 2012; Vestergard et al. 2008), be-
cause the Biolog EcoPlates were used to assess functional
diversity of the soil culturable bacterial community, while
the PCR-DGGE was focused on composition diversity of
the soil dominant bacterial community, and some bacterial
activity was susceptible prior to the community composi-
tion (Mijangos et al. 2009). Consequently, two microbial
methods used together provided a comprehensive under-
standing of the effects of BP on soil bacterial community
profiles. Considering the Biolog EcoPlates and PCR-
DGGE results for the UP treatment, functional diversity
of the soil culturable bacterial and composition diversity
of the soil dominant bacterial community both showed
little difference from CK in the whole incubation period;
for the BP treatment, composition diversity of the soil
dominant bacterial community showed little difference
from the control in the whole incubation period, whereas
activity and functional diversity of the soil culturable bac-
terial community were lower than the control in the early
incubation phase, and they recovered to the control level
in the 12
th
month of the incubation period.
Conclusions
During the 12 months of waterlogged incubation in paddy
soil, the biodegradation rate of BP was lower than that of
UP, the BP membrane remained more intact, and the soil
showed small influence on the structural integrity of the BP
membrane, which guaranteed the controlled release of nutri-
ents through the coating membrane. BP membranes, as well as
UP ones, indicated little impacts on composition diversity of
the soil dominant bacterial community. BP membranes sup-
pressed activity and functional diversity of the soil culturable
bacterial community at the early incubation phase but gradu-
ally recovered both in the 12
th
month, while UP showed no
significant negative effects on them over the whole incubation
period. Hence, waterborne polyacrylate materials showed
negligible harm to soil bacterial community profiles and were
environmentally friendly, and the biochar-modified mem-
brane not only improved the quality of CRF products but also
provided an alternative option for the utilization of crop
residues.
Tabl e 4 Composition diversity characterization of bacterial community in paddy soil using indexes of OTU richness (S), Shannon-Wiener index (H′),
and evenness (E)
OTU richness (S) Shannon-Wiener index (H′) Evenness (E)
Materials Sampling position Incubation period (month) Incubation period (month) Incubation period (month)
2 61226122612
CK Top layer 72ab 57a 47a 3.83a 3.49a 3.30a 0.90a 0.86a 0.86a
Bottom layer 73a 57a 49a 3.77a 3.61a 3.32a 0.88a 0.89a 0.85a
UP Top layer 74a 60a 48a 3.88a 3.39a 3.36a 0.90a 0.83a 0.87a
Bottom layer 72ab 61a 49a 3.78a 3.46a 3.40a 0.89a 0.84a 0.87a
BP Top layer 69b 59a 47a 2.46a 3.45a 3.28a 0.86a 0.84a 0.85a
Bottom layer 73ab 59a 49a 3.69a 3.55a 3.28a 0.88a 0.87a 0.84a
Means in a column followed by the same letter are not significantly different (P<0.05; Tukey test)
CK soil without polyacrylate membrane, UP soil with unmodified waterborne polyacrylate membrane, BP soil with biochar-modified polyacrylate
membrane, top layer the contacted soil layer (soil with 0–2-cm distance from the buried membrane surface), bottom layer the noncontact soil layer (soil
with 8–10-cm distance from the buried membrane surface)
Environ Sci Pollut Res
Acknowledgments This study was financially supported by National
12
th
Five-Year Science and Technology Supporting Program
(2011BAD11B01-02), R & D Projects from the Chinese Academy of
Sciences, and Special Fund for Agro-scientific Research in the Public
Interest (201303103). We would like to thank Prof. Xie Zubing for pro-
viding wheat-based biochar and Prof. Chu Haiyan for assisting with PCR-
DGGE.
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