Efficiency of phenol biodegradation by planktonic Pseudomonas pseudoalcaligenes (a constructed wetland isolate) vs. root and gravel biofilm.

This article appeared in a journal published by Elsevier. The attached
copy is furnished to the author for internal non-commercial research
and education use, including for instruction at the authors institution
and sharing with colleagues.
Other uses, including reproduction and distribution, or selling or
licensing copies, or posting to personal, institutional or third party
websites are prohibited.
In most cases authors are permitted to post their version of the
article (e.g. in Word or Tex form) to their personal website or
institutional repository. Authors requiring further information
regarding Elsevier’s archiving and manuscript policies are
encouraged to visit:
http://www.elsevier.com/copyright

Author's personal copy
Efficiency of phenol biodegradation by planktonic
Pseudomonas pseudoalcaligenes (a constructed wetland isolate)
vs. root and gravel biofilm
Eyal Kurzbauma, Felix Kirzhner a, Shlomo Sela b, Yoram Zimmels a,2, Robert Armon a,*,1
a Faculty of Civil & Environmental Engineering, Division of Environmental, Water & Agricultural Engineering,
Technion-Israel Institute of Technology, Haifa 32000, Israel
bDepartment of Food Science, The Volcani Center, Agricultural Research Organization, Bet Dagan, Israel
a r t i c l e i n f o
Article history:
Received 31 March 2010
Received in revised form
10 June 2010
Accepted 8 July 2010
Available online 16 July 2010
Keywords:
Pseudomonas pseudoalcaligenes
Rhizosphere
Phenol
Biodegradation
Biofilm
a b s t r a c t
In the last two decades, constructed wetland systems gained increasing interest in
wastewater treatment and as such have been intensively studied around the world. While
most of the studies showed excellent removal of various pollutants, the exact contribution,
in kinetic terms, of its particular components (such as: root, gravel and water) combined
with bacteria is almost nonexistent.
In thepresent study, a phenol degrader bacterium identified as Pseudomonas pseudoalcaligenes
was isolated from a constructed wetland, and used in an experimental set-up containing:
plants and gravel. Phenol removal rate by planktonic andbiofilmbacteria (on sterile Zeamays
roots and gravel surfaces) was studied. Specific phenol removal rates revealed significant
advantage of planktonic cells (1.04 � 10�9 mg phenol/CFU/h) compared to root and gravel
biofilms: 4.59 � 10�11e2.04 � 10�10 and 8.04 � 10�11e4.39 � 10�10 (mg phenol/CFU/h),
respectively.
In batch cultures, phenol biodegradation kinetic parameters were determined by biomass
growth rates and phenol removal as a function of time. Based on Haldane equation, kinetic
constants such as mmax ¼ 1.15/h, Ks ¼ 35.4 mg/L and Ki ¼ 198.6 mg/L fit well phenol removal
by P. pseudoalcaligenes.
Although P. pseudoalcaligenes planktonic cells showed the highest phenol removal rate, in
constructed wetland systems and especially in those with sub-surface flow, it is expected
that surface associated microorganisms (biofilms) will provide a much higher contribution
in phenol and other organics removal, due to greater bacterial biomass.
Factors affecting the performance of planktonic vs. biofilm bacteria in sub-surface flow
constructed wetlands are further discussed.
ª 2010 Elsevier Ltd. All rights reserved.
1. Introduction
Constructed wetland (CW) is an artificial marsh or swamp,
created for anthropogenic discharge treatment (such as
wastewater, agricultural and industrial effluents, and
stormwater runoff). CW-s are highly complex systems that
transform and remove various contaminants from waste-
water flow by different mechanisms (physical, chemical and
* Corresponding author. Tel.: þ972 48292377; fax: þ972 48293309.
E-mail address: cvrrobi@tx.technion.ac.il (R. Armon).
1 Member of Grand Water Research Institute, Technion, Haifa, Israel.
2 We would like to dedicate this article to Prof. Zimmels who prematurely passed away.
Avai lab le a t www.sc iencedi rec t .com
journa l homepage : www.e lsev ie r . com/ loca te /wat res
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 1
0043-1354/$ e see front matter ª 2010 Elsevier Ltd. All rights reserved.
doi:10.1016/j.watres.2010.07.020

Author's personal copy
biological) that occur sequentially or simultaneously (Kadlec
and Knight, 1996).
CW is basically a basin with a submerged porous bed layer
that holds different plants flooded with various effluents.
Porous bed (i.e. gravel) and vegetation that operate in a CW
provide surfaces upon which microorganisms can grow as
biofilms. The support porous beds act as a filter for suspended
solids through sedimentation processes due to the reduction
in stream velocity. In addition, plants due to their massive
roots surfaces are capable to remove some of the nutrients
(particularly phosphate and nitrogen), heavymetals and some
recalcitrant organics present in effluents (Kadlec and Knight,
1996; Vymazal, 2005). Microorganisms present in CW play
a key role in organic carbon removal. In a sub-surface flow
system, these microorganisms are roughly present in two
states: planktonic and sessile (biofilm). While planktonic
bacteria are continuously moving within the water system,
the biofilms are either attached onto plant roots or gravel bed
living in a customized microniche and complex microbial
community with homeostasis, metabolic cooperativity and
basic circulation (Costerton et al., 1995; Kadlec and Knight,
1996; Danhorn and Fuqua, 2007). Each biofilm bacterium
lives in a complex microbial community that has primitive
homeostasis, a primitive circulatory system, and metabolic
cooperativity, and each of these sessile cells reacts to its
special environment so that it differs fundamentally from
a planktonic cell of the same species.
More detailed characteristics of the various system
components and their environmental impacts were excel-
lently reviewed elsewhere (Kadlec and Knight, 1996;
Stottmeister et al., 2003; Vymazal, 2005).
In the recent years, wastewater treatment through CW
systems gained much interest and was studied intensively.
Nevertheless, most of the research on CW systems is mainly
descriptive based on engineering aspects, and less on the
existent basic interactions between system’s components. In
particular, there is lack of knowledge related to biochemical
interactions between bacterial population and the three
principal constituents: effluent water, plants and porous
media.
Beside their role as attachment surfaces, some researchers
squabble that plant roots provide a beneficial habitat for root-
attached microorganisms as a result of their biological nature
beside exudates such as: amino acids, simple sugars, complex
carbohydrates and oxygen (Gersberg et al., 1986; Brix, 1997;
Stottmeister et al., 2003). However, to date no experimental
data was presented regarding the beneficial habitat for
microorganisms provided by plant roots compared with inert
surfaces such as gravel bed in a CW environment. Further-
more, it is not clear whether the “rhizosphere effect” do
contribute significantly as a result of the continuous flow
around roots, resulting in wash up and dilution of root
exudates. Consequently, in a CW system the benefit of
plantemicrobial association in biodegradation acceleration
compared to inert surfaceemicrobial association (e.g. gravel)
is still unclear. On the other hand, macrophytes overall posi-
tive impact on pollutant removal processes in CW systems is
well documented (Tanner, 1996; Brix, 1997; Brisson and
Chazarenc, 2009; Zimmels et al., 2008). In this context, it is
not clear whether the reported positive effect is due to
increased surface area or plants provision of nutrients bene-
ficial to microorganisms metabolism.
A small number of studies have compared the character-
istics of different bacterial population present in CW systems
such as: the influence of plants’ presence onmicrobial density
and activity (Gagnon et al., 2007), activity of stem-attached
biofilm on emergent plant (Pollard et al., 1995), rhizosphere
and suspended populations biodegradation capabilities
(Toyama et al., 2006), and comparison of real plants, plastic
plants, absence of plants and bacterial communities on their
effect on water treatment in a CW mesocosms (Collins et al.,
2004). Albeit most studies pointed on plant roots and porous
bed as important surfaces involved in bacterial colonization,
no direct comparison between the two surfaces had been
made.
In the present study, a CW system isolated bacterium
Pseudomonas pseudoalcaligenes was characterized for its phenol
biodegradation capabilities (as a sole carbon and energy
source) in batch kinetics cultures and in three forms: plank-
tonic (as suspension), root attached and gravel attached (as
biofilm) in controlled experimental flasks. Phenol as
a common model for aromatic pollutant was selected due to
its popularity and gained knowledge on its biodegradation
(van Schie and Young, 2000).
2. Materials and methods
2.1. Bacterial and hydroponic growth minimal medium
The basic growth media were based on half-strength Hoag-
land’s minimal medium (HMM) (Hoagland and Arnon, 1950).
Hoagland’s stock nutrient medium was prepared by mixing
three stock solutions with the following formula: Solution 1
(macroelements) (g/L): KNO3 10.2, Ca(NO3)2$4H2O 7.08,
NH4H2PO4 2.3, MgSO4$7H2O 4.9 in double distilled water;
Solution 2 e (g/250 mL) FeSO4$7H2O 1.9, EDTA-Na2 1.25; and
Solution 3 (microelements) (g/L): H3BO3 2.86, MnCl2$4H2O 1.81,
CuSO4$5H2O 0.08, H2MoO4$H2O 0.09, ZnSO4$7H2O 0.22. Final
Hoagland’s composition was made up of solution 1, 2 and 3
(ratio 1:0.006:0.01) and distilled water up to 1 L. pH was
adjusted to 6.5 � 0.1 with sterile NaOH (1 M).
2.2. Isolation of phenol degrader bacterium from CW
Common Reed (Phragmites australis) plants were removed
from a vertical sub-surface flow CW mesocosm, as already
described by Zimmels et al. (2008). The system was supple-
mented with diluted domestic sewage (1:20 v/v) for 6 months
prior to sampling. To separate rhizospheral bacterial pop-
ulation, 5 g of excised root sections were vortexed in 200 mL
sterile saline (0.85% NaCl) containing Tween 20 (0.1%) (Sigma,
Israel) for 60 min on a rotary shaker (MRC, TS-400) at 200 rpm
and ambient temperature (22 � 2 �C). The suspension was
pre-filtered through a sterile cheese-cloth to remove large
particles. The filtrate was centrifuged for 10 min (6000�g) and
the pellet washed (�3) with sterile saline. Finally, 50 mL of
resuspended pellet was introduced into sterile 300 mL flask
containing 100 mL sterile HMM supplemented with 20 mg/L
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 15022

Author's personal copy
phenol (>99.5% pure) (Merck, USA) as the sole carbon and
energy source.
The inoculated flasks were shaken (at 100 rpm) for one
month at 30 � 1 �C, supplemented daily with 20 mg/L phenol.
From previous experimental results phenol at this concen-
tration was biodegraded completely in 24 h (data not shown)
therefore daily phenol addition was required to maintain
a selective environment. Weekly, the growing suspended
bacterial cells were centrifuge washed with saline (�3) and
used as inoculum in a fresh HMM medium containing phenol
as mentioned above. After one month, five samples from the
culture were spread on solid HMM medium (containing 15 g/L
Bacto agar, Difco, USA) supplemented with 50 mg/L phenol
(phenol-HMM) and incubated at 36 � 1 �C for 48 h. Solid HMM
medium without phenol was used as control.
Following incubation, a number of isolated colonies were
further plated and transferred several times using solid
phenol-HMM medium. One colony which showed a relatively
rapid growth on the phenol-HMMmedium (but could not grow
on the HMM medium without phenol) was isolated and
further examined for its properties and identified using 16S
ribosomal RNA analysis.
2.3. Identification of the isolated bacteria by 16S
ribosomal RNA analysis
Chromosomal DNA was extracted from the isolated bacteria
using the alkali-lysis technique, as described by Hartas et al.
(1998). Bacteria-domain-targeted PCR primers were used to
amplify a conserved region of bacterial DNA coding for 16S
rDNA (1392 bp), as described previously (Amann et al., 1995).
The following primers, purchased from SigmaGenosys (Israel)
were used: 11F (50-GTTTGATCMTGGCTCAG-30) and 1392R
(50-ACGGGCGGTGTGTAC-30). PCR was performed in a Bio-
metra/Tgradient thermocycler. Reactions were carried out in
a final volume of 40 ml, with 0.5 pmol of each primer and 20 ml
of PCR reaction mastermix (Fermentas, Ontario, Canada). PCR
conditions were: 95 �C for 4 min followed by 36 cycles of 94 �C
for 30 s, 58 �C for 30 s, and 72 �C for 1.5min. The final cycle was
followed by extension at 72 �C for 10 min. PCR products were
visualized on agarose gel (1%) following the purification step,
and sent for sequence determination (Hy Laboratories, Reho-
vot, Israel). Sequence identity (624 bp) was determined with
the BLAST program (Altschul et al., 1997).
Maximum likelihood and DNA sequences were analyzed
and aligned using the alignment tools implemented in ARB
(http://www.arb-home.de). Neighbor-joining tree was con-
structed using maximum likelihood (PhyML) in Phylogeny.fr
(http://www.phylogeny.fr). The robustness of the tree
topology was verified through calculating bootstrap (100
replicates) values for the neighbor-joining tree.
2.4. Phenol degradation kinetics and analytical
concentration measurement
The isolated P. pseudoalcaligenes bacteria was grown tomid-log
phase in 200mLHMMsupplementedwith 100mg/L phenol for
4 h on a rotary shaker (150 rpm) at 30 � 1 �C. The culture was
then centrifuged (10 min at 6000�g) and washed (�3) with
sterile saline. In kinetic experiments, inoculum from phenol
acclimated culture was split into 400 mL triplicate flasks.
Triplicates flasks contained 100 mL HMM solution were sup-
plemented with initial phenol concentrations of 10, 20, 40, 80,
180, 260, and 525 mg/L. The biodegradation experiments were
performed at 30 �C under continuous shaking (150 rpm).
Bacterial growth was measured by optical density at
600 nm (OD600) using a spectrophotometer (Spectronic 20
Genesys, Spectronic Instruments, USA). Phenol concentra-
tions were measured periodically using 4-aminoantipyrine
colorimetric method (APHA, 1995) on supernatant of samples
centrifuged for 10 min at 6000�g. For each phenol concen-
tration specific growth rate (m) and effective biodegradation
rate (dS/dt) were calculated from growth kinetic slope.
2.5. Sterile hydroponics plant growth conditions
Corn (Zea mays) (Gedera Seeds Co., Gedera, Israel) seeds were
surface sterilized for 20 min in 70% ethanol, 60 min in 2%
sodium hypochlorite followed by another 10 min in 70%
ethanol and 60 min in 1% sodium hypochlorite. Sterilized
seeds were washed 5 times in sterile HMM and then trans-
ferred into a sterile hydroponic chamber as previously
described (Burdman et al., 1996) containing sterile HMM.
Seven days old germinated sterile seedlings (approximately
4 cm long roots) were transferred to 250 mL bottles containing
100 mL HMM. While the roots were submerged into HMM
medium, the shoot emerged through the bottlemouth sealing.
The sealing was made of few aluminum foil and parafilm
layers wrapped tightly around the shoot and the bottle’s
mouth to prevent contamination and gases exchange. Plant
roots were kept in darkness by bottle wrapping with
aluminum foil. Whole plants were maintained in a growth
chamber at 22 �C with 18-h light/6-h dark photoperiod for 1
month. HMM medium was replaced under sterile conditions
once a week in order to provide continuous nutrient supply.
2.6. Experimental setup
Thirty seven days old sterile hydroponic grown Z. mays plants
were used for inoculation procedure. Z. mays plant was
chosen due to former experience and our ability to grow it
under sterile hydroponic conditions and as a gnotobiotic
plant. The gravel stones were limestone rock cut into small
squares (1 � 1 cm) (Degania Psifas, Israel) in order to facilitate
microscopic visualization and provided improved attachment
surface area. Experimental gravel cubes were washed inten-
sively in distilled water and autoclaved three times with fresh
distilled water each time in order to clean the gravel surface
from any adsorbed materials.
P. pseudoalcaligenes was grown overnight under continuous
shake (100 rpm) on a rotary shaker (MRC, TS-400) at 30 �C in
liquid HMM medium supplemented with 50 mg/L phenol as
a sole carbon. Grown culture was concentrated and washed
(�3) with sterile saline (0.85% NaCl) at 6000�g for 10 min. An
experimental solution (100 mL HMM containing 10 mM
MgSO4$7H2O after Fuqua and Matthysse (2001)) was made in
order to facilitate bacterial attachment onto root and gravel
surfaces. The final concentration of P. pseudoalcaligenes in
attachment solution was 3.6 � 109 CFU/mL.
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 1 5023

Author's personal copy
Roots and gravel inoculations with bacteria were per-
formed by dipping sterile roots and gravel into the above
solution for 2 h. Then roots and gravel were rinsedwith sterile
running water for 10 s (five times) and further dipped in
200 mL sterile HMM for additional 5 min. Preliminary experi-
ments confirmed that these procedures did not affect the
roots.
In order to provide two different densities of attached
bacteria on gravel and roots this procedure was repeated
twice. The first pretreatment consisted of inoculated plants
and gravel cubes immersed in HMM containing 20 mg/L
phenol for five days. In parallel, the second pretreatment was
set to maintain additional inoculated plants and gravel cubes
in HMM supplemented with 20 mg/L phenol and 1 g/L sodium
citrate (Sigma) for five days in order to create a denser bacte-
rial attached population. Citrate bioavailability of the present
strain have a selective effect as well a much simpler substrate
to be utilized compared to phenol, therefore increasing
bacterial biomass. Finally, after five days the solutions were
replaced with fresh HMM containing 20 mg/L phenol to
remove planktonic (unattached) bacteria and to acclimate the
attached bacteria to phenol prior to biodegradation experi-
ment setup. On the sixth day, plants and gravel with the two
population densities were transferred to new experimental
bottles where biodegradation experiments were performed.
The experimental system contained sterile 250 mL Erlen-
meyer flasks filled with sterile 100 mL HMM supplemented
with 20 mg/L phenol. In each experimental flask one plant
(10 � 1 g) was placed straightforward with roots immersed in
HMM medium and shoot emergent through the flask’s mouth
sealing (hermetically sealed as already described). For exper-
iments with gravel-attached bacteria, ten gravel cubes
carrying attached bacteria (15 g) were axenically placed in the
experimental flask. All gravel and root surfaces were
submerged in HMM medium supplemented with 20 mg/L
phenol.
Planktonic cultures were treated in flasks containing
100 mL HMM supplemented with 20 mg/L phenol and inocu-
lated with phenol acclimated culture to reach a concentration
of 1 � 106 CFU/mL. All treatments (root and gravel attached
and planktonic bacteria) were done in triplicates.
All experimental flasks were incubated in a plant growth
chamber under mild agitation (25 rpm) at 22 � 1 �C to provide
continuous mixed solution during the experiments.
Supernatant phenol concentration was measured inter-
mittently following sample centrifugation (6000�g) for 10min.
Bacterial concentrations were measured at the end of exper-
iment and specific removal rate was calculated for each
treatment (mg phenol/CFU/h).
2.7. Bacterial enumeration
Bacteria enumeration was calculated on the whole flask
population basis in order to avoid the complexity and the
difficulty of comparing specific surface area of gravel and
roots. All other possible methods to compare attached bacte-
rial population density on gravel and roots surfaces were
found to have unacceptable drawbacks.
Rhizosphere-attached bacteria separation was done as
previously described (Cook et al., 2006) with minor
modifications. Each beaker was covered and sealed tightly
with a thick layer of sterile aluminum foil. Experimental set-
ups containing plants consisted only roots in sealed beakers
after shoots slicing. To each beaker 3 drops of sterile Tween 80
and 2 g of sterile glass beads (3 mm diameter) were added and
hand shaken vigorously for 5 min then shaken on a rotary
shaker (MRC, TS-400) at 250 rpm for additional 10 min.
Bacteriawere enumerated on two differentmedia: nutrient
agar (Difco, Israel) and solid phenol-HMM medium (experi-
mentally found to be optimal to grow phenol degrading
bacteria). CFUwere counted on each plate (triplicates), and the
total CFU for each beaker was calculated as follows: total CFU/
flask ¼ (CFU/mL) � (total volume of medium).
2.8. Visualization of root attached and gravel attached
bacteria with confocal laser scanning microscopy (CLSM)
Bacterial colonization onto roots and gravel was detected by
use of the fluorogenic dye 40,60-diamidino-2-phenylindole
(DAPI, 100 mg/mL) after experiment conclusion. Colonizing
bacteria were stained by soaking detached roots sections and
gravel in DAPI solution for 3 min following rinses (�3) with
sterile HMM.
Detection and imaging of rhizoplane and gravel surfaces
attached bacteria were performed by CLSM (model: CTR
5500CS microscope, Leica, Germany) equipped with detectors
and filter sets that can simultaneously monitor red, blue and
green fluorescence. The greeneyellow autofluorescence
exhibited by root epithelia was used to visualize root surface
(for further details see Assmus et al., 1995). Image combina-
tion and process were performed with the standard software
package provided by Leica (Leica application Suite 3.0).
Sterile roots and gravel (without bacteria) were stained as
described above and examined using the same procedures as
controls.
2.9. Statistical analysis
Analysis of variance (ANOVA) was performed with SPSS
version 17.0 (SPSS, Chicago, IL, USA) with mean differences
significant at p � 0.01. Tukey’s post-hoc tests were used for
multiple comparisons between groups. In order to determine
kinetics parameters of phenol biodegradation by isolated
bacterium, a non-linear regression Haldane Model was fitted
using the original data, applying POLYMATH 5.1 forWindowsª
(2004-Control Data Corporation).
3. Results
3.1. Characterization and identification of the isolated
bacteria by 16S ribosomal RNA analysis
The CW bacterial rhizospheric isolate is a Gram (�) rod shape
bacteria. Colonies’ appearance grown on nutrient agar is pale
beige, round, smooth, and shiny. Under anaerobic conditions
no growth was observed (data not shown).
Similarity search through 16S rRNA sequence using the
BLAST program indicated a close relationship to bacterial
species P. pseudoalcaligenes (accession number: DQ777729)
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 15024

Author's personal copy
at 98.5% identity. Neighbor-joining tree of the isolated
P. pseudoalcaligenes demonstrating the relationship of the
isolated strain to other Pseudomonas sp. is shown in Fig. 1.
The sequence data have been submitted to the GenBank
database under accession number GQ327971.
The isolate was capable to utilize phenol as a sole carbon
and energy source. When grown on solid HMM supplemented
with 50 mg/L phenol vs. nutrient agar the CFU numbers were
identical. Solid HMM without any carbon sources did not
support growth.
3.2. Biodegradation of phenol
In order to examine the phenol biodegradation characteris-
tics of the newly CW isolate, P. pseudoalcaligenes, a series of
kinetic experiments were performed with pure planktonic
cultures (Figs. 2 and 3). Phenol initial concentration ranged
from 10 to 525 mg/L and bacterial initial inocula were set to
0.025e0.030 (OD600). Typical kinetic patterns of bacterial
biomass accumulation against phenol removal were
observed with different experimental initial phenol concen-
trations. Fig. 2 summarizes the results obtained for initial
phenol concentrations of 10, 20, 40 and 80 mg/L which
showed complete phenol consumption in 2, 2.5, 3.5, and
4.75 h respectively. Following phenol complete consumption,
bacterial biomass concentration remained constant until
experiment termination.
Above 80 mg/L and up to 525 mg/L phenol concentrations,
bacterial inhibition was observed (Fig. 3). At these initial
concentrations a stationary phase was observed after primary
utilization of 100e150 mg/L phenol, with further negligible
phenol removal up to 21 h. Bacterial biomass growth halted in
parallel to phenol inhibition effect.
In relation to published data on phenol biodegradation, it
can be assumed that the newly isolated P. pseudoalcaligenes is
inhibited by the intermediate compound catechol at phenol
concentrations of >160 mg/L. Catechol is formed during
phenol aromatic ring oxidation through monohydroxylation
by a mono-oxygenase phenol hydroxylase enzyme at ortho
Fig. 1 e Neighbor-joining tree showing the phylogenetic position of the newly constructed wetland isolate and related
species of the genus Pseudomonas based on partial 16S rRNA gene sequences. The GenBank accession number for each
microorganism used in the analysis is shown after the species name. Bootstrap values (expressed as percentage of 100
replicates) are shown at the branch. Bar represents 0.1 nucleotide substitutions per position.
0 1 2 3 4 5
0
10
20
30
40
50
60
70
80
90
A
)L/g
m(l
o
nehP
Time (Hours)
10 mg/l
20 mg/l
40 mg/l
80 mg/l
control (80 mg/l)
without bacteria
0 2 4 6 8
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
B
D
.
O(
ss
a
m
oiB
006
)
Time (Hours)
10 mg/L
20 mg/L
40 mg/L
80 mg/L
control - without phenol
Fig. 2 e Phenol removal (A) and biomass formation (B) by
P. pseudoalcaligenes at initial concentrations of 10, 20, 40,
and 80 mg/L.
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 1 5025

Author's personal copy
position on the pre-existing hydroxyl group (Agarry et al.,
2008) and is known by its inhibitory nature when accumu-
lates in growth media (Ampe and Lindley, 1996). While the
intermediary compound inhibition in bacterial growth and
phenol consumption was observed in batch cultures which
contained initial phenol concentration of 180mg/L and higher,
batch cultures with initial concentration of 80 mg/L and lower
did not show this phenomenon. We assume that degradation
of 80 mg/L phenol or lower do not produce enough interme-
diary compounds that cause inhibition.
In general, phenol biodegradation by bacterial pure
cultures has been adequately described by substrate inhibi-
tion models (Kumaran and Paruchuri, 1997). Haldane’s model
is the most commonly used among a number of substrate
inhibition models proposed for determination of kinetic
parameters dealing with inhibitory substrates (Yang and
Humphrey, 1975). Specific growth rates and substrate initial
concentration were plotted (Fig. 4) and used in Haldane model
which takes the form:
m ¼ mmax=ððKS=SÞ þ 1þ ðS=KiÞÞ
where mmax is the maximum growth rate in the absence of
inhibition; S is the steady state substrate concentration
(mg/L); Ks is the saturation constant, the lowest substrate
concentration (mg/L) at which, in the absence of inhibition,
the specific growth rate is half the maximum growth rate
(mmax) and Ki is the inhibition constant, which numerically
equals the highest substrate concentration (mg/L) at which
the specific growth rate is equal to one half the maximum
specific growth rate (mmax). Fig. 4 shows that Haldane model
fits well our experimental data for each batch culture (at
different initial phenol concentration) (R2 ¼ 0.89). Kinetic
parameters obtained for P. pseudoalcaligenes grown on
phenol as a sole carbon source were: mmax ¼ 1.15/h,
Ks ¼ 35.4 mg/L and Ki ¼ 198.67 mg/L. A significant decrease
in mmax of the isolate was observed at phenol concentration
of 180 mg/L. This may be due to inhibitory (catabolic
repression) (Agarry et al., 2008) and lytic effect of phenol as
previously reported (Ruiz-Ordaz et al., 1998).
Effective biodegradation rate (dS/dt) was calculated to each
initial phenol concentration in order to evaluate the capability
of P. pseudoalcaligenes to remove phenol (Fig. 5). Interestingly,
maximal biodegradation rate was observed at 180 mg/L, while
for the same initial concentration a complete growth and
substrate utilization inhibition occurred after consuming
100 mg/L phenol (Fig. 3). This inhibition is probably as well
a result of intermediate compounds inhibition (Ampe and
Lindley, 1996).
P. pseudoalcaligenes kinetic parameters (mmax, Ks, and Ki)
were compared with other phenol degrading microorganisms
(based on published data) (Table 1). Nevertheless, comparison
is only an estimate, as various data were obtained under
different experimental conditions. In spite of this, the repor-
ted mmax for P. pseudoalcaligenes is the highest, while Ks and Ki
were similar to those already reported. All other experiments
were performed at 20 mg/L phenol, as sub-inhibitory and
common representative phenol concentration of effluents
feed of CW-s (Kadlec and Knight, 1996).
0 5 10 15 20
0
100
200
300
400
500
600
A
)L/g
m(l
o
nehP
Time (Hours)
180 mg/L
260 mg/L
525 mg/L
control (525 mg/l)
without bacteria
0 2 4 6 8 10 12 14 16 18 20 22
0.00
0.02
0.04
0.06
0.08
0.10
0.12
0.14
0.16B
D
.
O(
ss
a
m
oiB
006
)
Time (Hours)
180 mg/L
260 mg/L
525 mg/L
Fig. 3 e Phenol removal (A) and biomass formation (B) by
P. pseudoalcaligenes at phenol initial concentration of 180,
260, and 525 mg/L.
0 100 200 300 400 500 600 700
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
et
ar
ht
w
org
cificepS
-
(
h
1
-
)
Initial phenol concentration (mg/L)
Experimental
values
µ
Fig. 4 e Comparison of experimentally specific growth rate
(m)with thatof thepredictedbyHaldane’smodel. Thespecific
growth rate for each phenol initial concentration was
calculated from the growth kinetics of P. pseudoalcaligenes
grown on phenol as the sole carbon source. The line
representsthebestfit (R2[0.89) to theHaldanemodelusedto
describe substrate inhibition kinetics.
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 15026

Author's personal copy
When non-biological processes contribution by experi-
mental components (gravel, plant andwater) was evaluated, it
had been found that negative controls such as sterile solution
and sterile gravel did not alter phenol concentration due to
non-specific abiotic adsorption or volatilization. Sterile plant
control showed a minor reduction in phenol concentration
mostly due to passive adsorption (from 19.1 to 17.3 mg/L
during 50 h interval) (data not shown). These controls revealed
that nonbacterial processes are negligible factors in phenol
removal.
Asmentioned above, P. pseudoalcaligeneswas present in our
experimental set-up in two states: biofilm and planktonic.
Phenol biodegradation characteristics of these two forms
were examined. In addition, biofilm bacteria were examined
at two densities. Fig. 6 shows phenol removal by P. pseu-
doalcaligenes planktonic and biofilm cells after five days
pretreatment with 20 mg/L phenol. Planktonic bacteria
showed complete phenol removal in 4.33 h, while roots and
gravel biofilm bacteria showed complete phenol removal in
6.33 and 6.83 h, respectively.
Fig. 7 shows phenol removal by P. pseudoalcaligenes root and
gravel biofilm, pretreated with 20 mg/L phenol and 1 g/L
sodium citrate in HMM solution. Root and gravel biofilms
removed phenol completely in 4 h. The higher phenol removal
rate by biofilm cells in this case is mainly a direct result of
increased viable bacterial numbers as shown in Table 2. Pre-
exposure of surface associated bacteria in a richer organic
solution (citrate þ phenol) enabled formation of a denser
bacterial biofilm on gravel and root surfaces.
As each experimental set-up varied in relation to bacterial
numbers, it was required to normalize through specific
phenol removal rate calculation (mg phenol removed/CFU/h)
(Fig. 8). Planktonic cells (1.04 � 10�9 mg phenol/CFU/h)
revealed a significant higher specific phenol removal rate
( p < 0.01) compared to root and gravel biofilms:
4.59 � 10�11e2.04 � 10�10 and 8.04 � 10�11e4.39 � 10�10 (mg
phenol/CFU/h), respectively.
As shown in Table 2, gravel and root biofilms grown with
supplemental citrate revealed one order of magnitude higher
cells density compared to phenol alone. However, specific
removal rate of biofilms grown with supplemental citrate was
lower by half order of magnitude.
Finally, spatial distribution of P. pseudoalcaligenes biofilms
associated with roots and submerged gravel surfaces, were
observed with CLSM (Fig. 9aed). Fig. 9 reveals that biofilm
covering roots and gravel surfaces were scattered with occa-
sional clustering (Fig. 9a and d). The two pre-exposure
0 100 200 300 400 500 600 700
0
10
20
30
40
et
ar
n
oit
ad
arged
oibl
o
nehP
)h/L/g
m(
Phenol initial concentration (mg/L)
Fig. 5 e Phenol biodegradation rates (dS/dt, mg/L/h) as
a function of initial phenol concentration.
Table 1 e Comparison of kinetic constants (the parameters were obtained from the Haldane’s model) for phenol
biodegradation from various studies and the present one.
Microorganism mmax (1/h) Ks (mg/L) Ki (mg/L) Reference
Pseudomonas pseudoalcaligenes (CW isolate) 1.15 35.4 198.67 This study
Pseudomonas resinovorans 1.007 13 117.7 Dikshitulu et al. (1993)
Pseudomonas putida 0.569 18.5 99.4 Beyenal et al. (1997)
Pseudomonas fluorescens 0.618 71.4 241 Kumaran and Paruchuri (1997)
Bacillus circulans 0.27 1.75 1743 Vijayagopal and Viruthagiri (2005)
Rhodococcus sp. 0.33 0.32 1264 Straube et al. (1990)
Nocardia sp. 0.422 26.77 261.72 Vijayagopal and Viruthagiri (2005)
Acinetobacter calcoaceticus 0.542 36.2 145 Kumaran and Paruchuri (1997)
Candida tropicalis 0.375 3.7 848 Ruiz-Ordaz et al. (1998)
0 1 2 3 4 5 6 7
0
5
10
15
20
25
n
oit
art
nec
n
ocl
o
nehP
)L/g
m(
Time (hours)
gravel
root
planktonic
sterile (cont.)
Fig. 6 e Phenol removal by planktonic and root/gravel
biofilm of P. pseudoalcali genes, pre-exposed for five days to
20 mg/L phenol in HMM solution.
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 1 5027

Author's personal copy
treatments (phenol with or without citrate) did not show
visual colonization differences.
4. Discussion
The present study experimental results suggest that biofilm
and planktonic cells of P. pseudoalcaligenes have different
phenol removal efficiencies. Measuring specific phenol
removal rates it was found that planktonic cells were quan-
titatively 22 times higher in comparison with root and gravel
biofilms, while other processes (i.e. sterile medium, plant and
gravel) were negligible.
Citrate pretreatment (in addition to phenol) for 5 days
increased biofilm cells density on gravel and root surfaces
compared to phenol solely, though specific phenol removal
rate declined. van Loosdrecht et al. (1990) showed a similar
phenomenon, described by them as reduced phenol and
oxygen mass transfer through denser biofilms. This issue is
still controversial in the scientific literature. Some researchers
have pointed out that planktonic Pseudomonas sp. cells are
relatively more efficient in substrate removal compared to
biofilms (Barton et al., 1996; Heffernan et al., 2009) as well with
other bacterial species (Barton et al., 1996; Jeffrey and Paul,
1986). On the other hand, other studies showed that biofilm
cells could remove more efficiently different substrates
coupled with increased respiration rates compared to plank-
tonic cells (Fletcher, 1979; Ascon-Cabrera et al., 1995; Bester
et al., 2005), while other authors found no significant differ-
ence between the two states (Irriberry et al., 1990; Bright and
Fletcher, 1983). Generally, attachment affects microorgan-
isms (including bacteria) in terms of activity and other
parameters as already reviewed and discussed elsewhere
(van Loosdrecht et al., 1990).
Phenol is a low-molecular-weight substrate, therefore it is
likely to be removed rapidly by planktonic bacteria and its
availability for biofilms is often reduced substantially by
diffusion and mass transfer (Marshall, 1992). The lower
phenol specific removal rates by biofilms (attached cells)
obtained in this study can be justified by these effects.
Nevertheless, in CW systems and especially in those with
sub-surface flow, it is expected that biofilm microorganisms
will provide a much higher contribution in organics removal
compared to planktonic ones, due to their increased bacterial
biomass (Pollard et al., 1995; Polprasert and Khatiwada, 1998).
In a previous study, that analyzed the relative contribution of
different system components in phenol removal of a pilot
Table 2 e Bacterial concentration of planktonic, root biofilm, and gravel biofilm of P. pseudoalcaligenes, in the five
experimental setups.
Pre-exposure treatment Bacterial state CFU/flask S.D.
20 mg/L phenol in HMM Gravel attached 7.22 � 108 1.82 � 108
20 mg/L phenol in HMM Root attached 2.14 � 109 1.09 � 109
20 mg/L phenol þ 1 g/L sodium citrate in HMM Gravel attached 7.20 � 109 3.10 � 109
20 mg/L phenol þ 1 g/L sodium citrate in HMM Root attached 1.29 � 1010 5.40 � 109
20 mg/L phenol in HMM Planktonic 1.02 � 109 2.83 � 107
0 1 2 3 4 5
0
5
10
15
20
25
n
oit
art
nec
n
ocl
o
nehP
)L/g
m(
Time (hours)
gravel
root
planktonic
sterile (cont.)
Fig. 7 e Phenol removal as a function of time by
P. pseudoalcaligenes root and gravel biofilms (following five
days pre-exposure to 20 mg/L phenol and 1 g/L sodium
citrate in HMM solution) and compared to planktonic cells.
0.0
2.0x10
4.0x10
6.0x10
8.0x10
1.0x10
1.2x10
GPC
PPC
PPRP
GP
d cd
c
b
a
et
arl
a
v
o
merl
o
nehp
cificepS
)h/
UF
C/g
m(
Treatments
Fig. 8 e Box-and-whisker diagram representing specific
phenol removal rate results (mg phenol/CFU/h) of
planktonic and biofilm P. pseudoalcaligenes cells. Biofilm
cells include the two pre-exposure treatments: HMM
supplemented with 20 mg/L phenol (GP-gravel attached,
RP-root attached) and HMM supplemented with 20 mg/L
phenol and 1 g/L sodium citrate (PPC-root attached, GPC-
gravel attached). Letters beside each box represent Tukey’s
test results of significance differences ( p £ 0.01).
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 15028

Author's personal copy
scale CW, it was found that 95% of phenol removal could be
attributed to biodegradation by biofilms (on root and gravel)
and only 5% to planktonic population (Kurzbaum et al., in
press).
It iswidely accepted that plant root surface provides a highly
productive habitat increasing microorganisms’ biomass on
rhizoplane and in rhizosphere (Varma et al., 2004). In terrestrial
habitats, roots provide a nutrient-rich environment for micro-
organisms by release of many metabolites (Tsao, 2003),
including compounds that induce microbial genes involved in
pollutant degradation (Gilbert and Crowley, 1997; Casavant
et al., 2003) and acting as co-metabolites that facilitate degra-
dationof certain recalcitrantmaterials (Rentz et al., 2005). In the
present study it has been shown that root surface biofilm was
almost equally efficient to gravel biofilm in phenol specific
removal. A possible explanation to this similarity can be
attributed to roots adjacent water that may dilute roots
exudates in comparison to available high phenol concentra-
tions, making their role negligible. Thus, in our designed
hydroponic grown plants (or possibly in a real CW system)
rhizosphere advantage usually attributed to exudates effect in
terrestrial habitat is less pronounced in the aquatic one.
Micrographs of attached bacteria on hair, tips and main
root grownhydroponically, showed a scattered pattern (Fig. 9).
This pattern is not consistent with data reported for other
Pseudomonas species that showed mainly bacterial coloniza-
tion at borders of adjacent plant root cells in soil (Hansen
et al., 1997; Ramos et al., 2000; Pliego et al., 2008). This differ-
ence can be related to growing environment dissimilarity
between the two experimental systems.
Further studies should focus on correlation between bio-
film thickness attached onto gravel and plant roots and
certain nutrients utilization rate. Better understanding of
processes and mechanisms that dominate water purification
Fig. 9 e CLSM in situ visualization of DAPI stained P. pseudoalcaligenes bacteria colonizing root and gravel surfaces (a) gravel
surface; (b) main root (inset-sterile root cells); (c) root tip; (d) root hair.
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 1 5029

Author's personal copy
through the gravel attached, root attached and planktonic
bacteria is essential in order to improve the purification
performance of CW systems. So far, most studies dealing with
CW systems, reported mainly empirical monitoring of reduc-
tion properties of different parameters in CW systems, and
descriptive-engineering aspects oriented to system perfor-
mance improvement. The current study dealtwith the specific
components contribution to the total removal of phenol, as
a model contaminant. Further study on interactions between
plants, microorganisms and support media are likely to
contribute to our knowledge on synergy/antagonism between
these components.
5. Conclusions
1. Planktonic cells showed a superior phenol removal rate
compare to root and gravel biofilms. Nevertheless, in sub-
surface flow CW systems, biofilm bacteria are expected to
be the major contributors to phenol removal based on its
significant larger cell number density compared to plank-
tonic population. In addition, the specificmaximumgrowth
rate (mmax) of P. pseudoalcaligenes on phenol was found to be
the highest compared to other microorganisms (Table 1).
Further studies will be performed in order to find out if this
event is universal or a singularity of P. pseudoalcaligenes.
2. Interestingly, phenol removal by the two biofilms (roots and
gravel) was almost equally efficient in spite of roots
exudates that were expected to provide a significant
advantage to its biofilm.
Acknowledgements
This research was partially supported by the Faculty of Civil
and Environmental Engineering and Grand Water Research
Institute at Technion. The authors would like to thank to
Dr. S. Avrahami for her helpful advice during construction of
the phylogenetic tree and to Lora Parahovnik for her labora-
tory assistance (Technion-Israel Institute of Technology).
Nidal Masalha (The Galilee Society, Israel) is acknowledged
for his help in kinetics evaluation.
r e f e r e n c e s
Agarry, S.E., Durojaiye, A.O., Solomon, B.O., 2008. Microbial
degradation of phenols: a review. International Journal of
Environment and Pollution 32, 12e28.
Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J., Zhang, Z.,
Miller, W., Lipman, D.J., 1997. Gapped BLAST and PSI-BLAST:
a new generation of protein database search programs.
Nucleic Acids Research 25, 3389e3402.
Amann, R.I., Ludwig, W., Schleifer, K.H., 1995. Phylogenetic
identification and in situ detection of individual microbial
cells without cultivation. Microbiological Reviews 59, 143e169.
Ampe, F., Lindley, N.D., 1996. Flux limitations in the ortho
pathway of benzoate degradation of Alcaligenes eutrophus:
metabolite overflow and induction of the meta pathway at
high substrate concentrations. Microbiology 142, 1807e1817.
APHA, 1995. Standard Methods for the Examination of Water and
Wastewater, 19th ed. American Public Health Association,
American Water Works Association, Water Environment
Federation, Washington, DC.
Ascon-Cabrera, M.A., Ascon-Reyes, D.B., Lebeault, J.M., 1995.
Degradation activity of adhered and suspended Pseudomonas
cells cultured on 2,4,6-trichlorophenol, measured by indirect
conductimetry. Journal of Applied Bacteriology 79, 617e624.
Assmus, B., Hutzler, P., Kirchhof, G., Amann, R., Lawrence, J.R.,
Hartmann, A., 1995. In situ localization of Azospirillum
brasilense in the rhizosphere of wheat with fluorescently
labeled, rRNA-targeted oligonucleotide probes and scanning
confocal laser microscopy. Applied and Environmental
Microbiology 61, 1013e1019.
Barton, A.J., Sagers, R.D., Pitt, W.G., 1996. Measurement of
bacterial growth rates on polymers. Journal of Biomedical
Materials Research Part A 32 (2), 271e278.
Bester, E., Wolfaardt, G., Joubert, L., Garny, K., Saftic, S., 2005.
Planktonic-cell yield of a Pseudomonad biofilm. Applied and
Environmental Microbiology 71, 7792e7798.
Beyenal, H., Seker, S., Tanyolac, A., Salih, B., 1997. Diffusion
coefficients of phenol and oxygen in a biofilm of Pseudomonas
putida. AIChE Journal 43, 243e250.
Bright, J.J., Fletcher, M., 1983. Amino acid assimilation and
electron transport system activity in attached and free-living
marine bacteria. Applied and Environmental Microbiology 45,
818e825.
Brisson, J., Chazarenc, F., 2009. Maximizing pollutant removal in
constructed wetlands: should we pay more attention to
macrophyte species selection? Science of the Total
Environment 407, 3923e3930.
Brix, H., 1997. Do macrophytes play a role in constructed
treatment wetland? Water Science and Technology 35, 7e11.
Burdman, S., Volpin, H., Kigel, J., Kapulnik, Y., Okon, Y., 1996.
Promotion of nod gene inducers andnodulation in commonbean
(Phaseolus vulgaris) roots inoculated withAzospirillum brasilense.
Applied and Environmental Microbiology 62 (8), 3030e3033.
Casavant, N.C., Thompson, D., Beattie, G.A., Phillips, G.J.,
Halverson, L.J., 2003. Use of a site-specific recombination-
based biosensor for detecting bioavailable toluene and related
compounds on roots. Environmental Microbiology 5, 238e249.
Collins, B.S., McArthur, J.V., Sharitz, R.R., 2004. Plant effects on
microbial assemblages and remediation of acidic coal pile
runoff in mesocosm treatment wetlands. Ecological
Engineering 23, 107e115.
Cook, K.L., Garland, J.L., Layton, A.C., Dionisi, H.M., Levine, L.H.,
Sayler, G.S., 2006. Effect of microbial species richness on
community stability and community function in a model
plant-based wastewater processing system. Microbial Ecology
52, 725e737.
Costerton, J.W., Lewandowski, Z., Caldwell, D.E., Korber, D.R.,
Lappin-Scott, H.M., 1995. Microbial biofilms. Annual Review of
Microbiology 49, 711e745.
Danhorn, T., Fuqua, C., 2007. Biofilm formationbyplant-associated
bacteria. Annual Review of Microbiology 61, 401e442.
Dikshitulu, S., Baltzis, B.C., Lewandowski, G.A., Pavlou, S., 1993.
Competition between two microbial populations in
a sequencing fed-batch reactor theory, experimental
verification, and implications for waste treatment
applications. Biotechnology and Bioengineering 42, 643e656.
Fletcher, M., 1979. A microautoradiographic study of the activity
of attached and free-living bacteria. Archives of Microbiology
122, 271e274.
Fuqua, C., Matthysse, A.G., 2001. Methods for studying bacterial
biofilms associated with plants. Methods in Enzymology 337,
3e18.
Gagnon, V., Chazarenc, F., Comeau, Y., Brisson, J., 2007. Influence
of macrophyte species on microbial density and activity in
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 15030

Author's personal copy
constructed wetlands. Water Science and Technology 56 (3),
249e254.
Gersberg, R.M., Elkins, B.V., Lyon, S.R., Goldman, C.R., 1986. Role
of aquatic plants in wastewater treatment by artificial
wetlands. Water Research 20, 363e368.
Gilbert, E.S., Crowley, D.E., 1997. Plant compounds that induce
polychlorinated biphenyl biodegradation by Arthrobacter sp.
strain B1B. Applied and Environmental Microbiology 63,
1933e1938.
Hansen, M., Kragelund, L., Nybroe, O., Sørensen, J., 1997. Early
colonization of barley roots by Pseudomonas fluorescens studied
by immunofluorescence technique and confocal laser
scanning microscopy. FEMS Microbiology Ecology 23, 353e360.
Hartas, J., Hibble, M., Sriprakash, K.S., 1998. Simplification of
a locus-specific DNA typing method (VirTyping) for
Streptococcus pyogenes. Journal of Clinical Microbiology 36,
1428e1429.
Heffernan, B., Murphy, C.D., Casey, E., 2009. Comparison of
planktonic and biofilm cultures of Pseudomonas fluorescens
DSM 8341 cells grown on fluoroacetate. Applied and
Environmental Microbiology 75, 2899e2907.
Hoagland, D.R., Arnon, D.I., 1950. The WatereCulture Method for
Growing Plants without Soil. Circular 347. Agricultural
Experiment Station, University of California, Berkeley.
Irriberry, J., Unanue, M., Ayo, B., Barcina, I., Egea, L., 1990.
Bacterial production and growth rate estimation from [3H]
thymidine incorporation for attached and free-living bacteria
in aquatic systems. Applied and Environmental Microbiology
56, 483e487.
Jeffrey, W.H., Paul, J.H., 1986. Activity of an attached and
free-living Vibrio sp. as measured by thymidine incorporation,
p-iodonitrotetrazolium reduction, and ATP/ADP ratios.
Applied and Environmental Microbiology 51, 150e156.
Kadlec, R., Knight, R., 1996. Treatment Wetlands. Lewis
Publishers, Chelsea, MI, USA.
Kumaran, P., Paruchuri, Y.L., 1997. Kinetics of phenol
biotransformation. Water Research 31, 11e22.
Kurzbaum, E., Zimmels, Y., Kirzhner, F., Armon, R. Removal of
phenol in a constructed wetland system and the relative
contribution of plant roots, microbial activity and porous bed.
Water Science and Technology, in press.
van Loosdrecht, M.C., Lyklema, J., Norde, W., Zehnder, A.J., 1990.
Influence of interfaces on microbial activity. Microbiology and
Molecular Biology Reviews 54, 75e87.
Marshall, K.C., 1992. Planktonic versus sessile life of prokaryotes.
In: Balows, A., Truper, H.G., Dworkin, M., Harder, W.,
Schleifer, K.H. (Eds.), The Prokaryotes: A Handbook on the
Biology of Bacteria: Ecophysiology, Isolation, Identification,
Applications. Springer-Verlag, New York, pp. 262e275.
Pliego, C., de Weert, S., Lamers, G., de Vicente, A., Bloemberg, G.,
CazorlaRamos, C., 2008. Two similar enhanced root-colonizing
Pseudomonas strains differ largely in their colonization
strategies of avocado roots and Rosellinia necatrix hyphae.
Environmental Microbiology 10, 3295e3304.
Pollard, P.C., Flood, J.A., Ashbolt, N.J., 1995. The direct
measurement of bacterial growth in biofilms of emergent
plants (Schoenoplectus) of an artificial wetland. Water Science
and Technology 32, 251e256.
Polprasert, C., Khatiwada, N.R., 1998. An integrated kinetic model
for water hyacinth ponds used for wastewater treatment.
Water Research 32 (1), 179e185.
Ramos, C., Mølbak, L., Molin, S., 2000. Bacterial activity in the
rhizosphere analyzed at the single-cell level by monitoring
ribosome contents and synthesis rates. Applied and
Environmental Microbiology 66, 801e809.
Rentz, J.A., Alvarez, P.J.J., Schnoor, J.L., 2005. Benzo[a]pyrene
co-metabolism in the presence of plant root extracts and
exudates: implications for phytoremediation. Environmental
Pollution 136, 477e484.
Ruiz-Ordaz, N., Manzano, W.H., Lagu´nez, J.C.R., Urbina, E.C.,
Mayer, J.G., 1998. Growth kinetic model that describes the
inhibitory and lytic effects of phenol on Candida tropicalis
yeast. Biotechnology Progress 14, 966e969.
van Schie, P.M., Young, L.Y., 2000. Biodegradation of phenol:
mechanisms and applications. Bioremediation Journal 4,
1e18.
Stottmeister, U., Wiesner, A., Kuschk, P., Kappelmeyer, M.,
Kaster, M., 2003. Effects of plants and microorganisms in
constructed wetlands for wastewater treatment.
Biotechnology Advances 22, 93e117.
Straube, G., Hensel, J., Niedan, C., Straube, E., 1990. Kinetic studies
of phenol degradation by Rhodococcus sp. P1 I. Batch
cultivation. Antonie Van Leeuwenhoek 57 (1), 29e32.
Tanner, C.C., 1996. Plants for constructed wetland treatment
systems e a comparison of the growth and nutrient uptake
characteristics of eight emergent species. Ecological
Engineering 7, 59e83.
Toyama, T., Yu, N., Kumada, H., Sei, K., Ike, M., Fujita, M., 2006.
Accelerated aromatic compounds degradation in aquatic
environment by use of interaction between Spirodela polyrrhiza
and bacteria in its rhizosphere. Journal of Bioscience and
Bioengineering 101, 346e353.
Tsao, D.T., 2003. Phytoremediation. In: Advances in Biochemical
Engineering/Biotechnology, vol. 78. Springer, Berlin.
Varma, A., Abbott, L., Werner, D., Hampp, R., 2004. Plant Surface
Microbiology. Springer, Germany.
Vijayagopal, V., Viruthagiri, T., 2005. Kinetics of biodegradation of
phenol using mixed culture isolated from mangrove soil.
Pollution Research 24, 157e162.
Vymazal, J., 2005. Horizontal sub-surface flow and hybrid
constructed wetlands systems for wastewater treatment.
Ecological Engineering 25, 478e490.
Yang, R.D., Humphrey, A.E., 1975. Dynamic and steady state
studies of phenol biodegradation in pure and mixed cultures.
Biotechnology and Bioengineering 17, 1211e1235.
Zimmels, Y., Kirzhner, F., Schreiber, J., 2008. Removal of high
organic loads from winery wastewater by aquatic plants.
Water Environment Research 80, 806e822.
wat e r r e s e a r c h 4 4 ( 2 0 1 0 ) 5 0 2 1e5 0 3 1 5031