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Structure of nitrifying biofilms in a high-rate trickling filterdesigned for potable water pre-treatment

Ben van den Akker a,*, Mike Holmes b, Peter Pearce c, Nancy J. Cromar a,Howard J. Fallowfield a

aHealth and Environment Group, School of the Environment, Flinders University, Bedford Park, South Australia 5042, AustraliabUnited Water International, 180 Greenhill Road, Parkside, South Australia 5063, AustraliacThames Water Research and Development, Reading Sewage Treatment Works, Island Road, Reading RG2 0RP, UK

a r t i c l e i n f o

Article history:

Received 15 December 2010

Received in revised form

2 March 2011

Accepted 11 April 2011

Available online 15 April 2011

Keywords:

Nitrification

Trickling filter

FISH

Scanning electron microscopy

Organic carbon

Biological water treatment

Carbohydrate

Protein

Heterotrophs

Ammonia

Multi-species biofilm

a b s t r a c t

This study examined the composition and structure of nitrifying biofilms sampled from

a high-rate nitrifying trickling filter which was designed to pre-treat raw surface water for

potable supply. The filter was operated under a range of feed water ammonia and organic

carbon concentrations that mimicked the raw water quality of poorly protected catch-

ments. The biofilm structure was examined using a combination of fluorescence in situ

hybridisation and scanning electron microscopy. Biopolymers (carbohydrate and protein)

were also measured. When the filter was operated under low organic loads, nitrifiers were

abundant, representing the majority of microorganisms present. Uniquely, the study

identified not only Nitrospira but also the less common Nitrobacter. Small increases in

organic carbon promoted the rapid growth of filamentous heterotrophs, as well as the

production of large amounts of polysaccharide. Stratification of nitrifiers and heterotrophs,

and high polysaccharide were observed at all filter bed depths, which coincided with the

impediment of nitrification throughout most of the filter bed. Observations presented here

specifically linked biofilm structure with filter functionality, physically validating previous

empirical modelling hypotheses regarding competitive interactions between autotrophic

and heterotrophic bacteria in biofilms.

ª 2011 Elsevier Ltd. All rights reserved.

1. Introduction

Uncontrolled variability in the concentration of ammonia in

feed waters during potable water treatment presents both

difficult and expensive challenges for maintaining effective

process control of chlorine disinfection. The conventional

approach to managing the level of influent ammonia in this

situation is pre-chlorination. This process, however, has

a number of recognized disadvantages, some of which

include: high chlorine dose requirements under high influent

ammonia loads; difficult process control where feed water

ammonia concentrations are unstable; the production of

harmful disinfection by-products; formation of chlorine-

related tastes and odours; and high process costs associated

with high chlorine demand.

Due to the deterioration of raw water quality worldwide

and increased demands for human consumption, water

treatment processes need to be improved to make the water

* Corresponding author. Present address: Water Research Centre, School of Civil and Environmental Engineering, The University of NewSouth Wales, Sydney 2052, Australia.

E-mail address: b.vandenakker@unsw.edu.au (B. van den Akker).

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journa l homepage : www.e lsev ie r . com/ loca te /wat res

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0043-1354/$ e see front matter ª 2011 Elsevier Ltd. All rights reserved.doi:10.1016/j.watres.2011.04.017

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safe and aesthetically pleasing, without the use of large

chlorine doses. This view was recognized in the 1980’s when

the U.S. Environmental Protection Agency took action to

regulate chlorination by-products (Sayre, 1988; Rittmann and

Huck, 1989). Early review papers by Rittmann and colleagues

in the 1980’s found biological treatment to be a cost-effective

and necessary solution to remove biodegradable contami-

nants such as ammonia, so that the formation of disinfection

by-product can be minimized (Rittmann and Snoeyink, 1984;

Rittmann and Huck, 1989). Although biological treatment of

public water supplies is not widespread practice throughout

the world, it has been relatively common in Europe since the

1970’s, mainly due to poorer surface water quality (Rittmann

and Huck, 1989).

There are a variety of approaches to biologically treat

drinking water which include: slow sand filtration, bank

filtration, biologically active granular activated carbon,

submerged aerated filters, unsubmerged filters, and fluidized

beds (Rittmann and Huck, 1989; Ahmad, 2005). Nitrifying

trickling filters (NTFs), which are conventionally used for

wastewater treatment, are another form of biological treat-

ment that have been successfully applied for ammonia

removal as well as the removal of manganese and iron from

ground and surface waters (e.g. Tekerlekopoulou and

Vayenas, 2007, 2008, van den Akker et al., 2008; De Vet et al.,

2009; Tekerlekopoulou et al., 2010). Interest in NTFs is likely

due to the well-established advantages of these systems such

as simple design and operation, low capital and operating cost

and a low requirement for energy and maintenance inputs.

Todate,most of our current understandingofNTFshas come

from research on wastewater applications, which are largely

based on one dimensional monitoring and manipulation of

traditional bulk physicalechemical parameters; with limited

scope for characterising filter microbiology. A key observation

from these traditional macroscale studies is that ammonia

removal (nitrification) is increasingly suppressedwith increased

organic carbon loading (see Boller and Gujer, 1986; Parker and

Richards, 1986; Pearce and Williams, 1999; Pearce and

Edwards, 2010). This suppression of nitrification has been

attributed to a response by faster growing heterotrophic micro-

organisms to increasedorganic carbonsubstrate availability and

subsequent out-competition of slower growing, autotrophic

nitrifiers. The presence of a rapidly respiring heterotrophic bio-

film layer in between the nitrifying populations and the

oxygenated bulk liquid film restricts or prevents the diffusion of

dissolved oxygen (DO) to the underlying nitrifiers, resulting in

lower rates of ammonia oxidation. Satoh et al. (2000) and Zhang

et al. (1994) have demonstrated this heterotrophic overgrowth of

nitrifying biofilms at the laboratory-scale by exposing coupon-

attached biofilms to varying C/N substrate ratios. Without

complementary researchat the full-scalehowever, it canonlybe

assumed that this pattern reflects the functional biofilm

processes of larger-scale percolating or flooded filters.

The impact of oxygen transfer as a key mechanism influ-

encing NTF performance has been described by Pearce and

colleagues with respect to increasing irrigation velocity driving

increased oxygen transfer for satisfying the metabolic require-

ments of both heterotrophs and autotrophs (Pearce and

Williams, 1999; Pearce and Edwards, 2010). The authors devel-

opedmodels for predicting effluent ammonia concentrations in

wastewater NTFsdboth stone and plastic mediadwith respect

to five day biological oxygen demand (BOD5), ammonia and

hydraulic loading (Pearce and Williams, 1999; Pearce and

Edwards, 2010). However, since these models were empirically-

based, heterotropheautotroph competition was only proposed

as one potential factor affecting nitrification performance in the

presence of organic carbon substrates; with this hypothesis

remaining un-validated.

In order to better design and optimise nitrification processes,

it is necessary to first identify the key organisms involved in this

microbial-driven processes and then to understand the func-

tional relationships between the microbiology and the process

performance.Understandingmicrobial functions and thenusing

this information in an appliedmanner should be fundamental to

NTFoperations,yet this is rarely thecase inpractice.Todate,very

few studies have characterised the microbiology of wastewater

NTFs (e.g. Persson et al., 2002; Biesterfeld et al., 2003; Lydmark

et al., 2006; Datta et al., 2011). Moreover, even fewer studies

have characterized and contrasted the physicalechemical

performance andmicrobiology of high-rate, potable water NTFs

e particularly under low ammonia and organic carbon substrate

conditions that are commonly encountered in poorly protected

catchments throughout the world.

Theaimof this studywas to characterise themicrobiological

performance of a potablewater NTF operating under a range of

controlled ammonia and soluble BOD5 (sBOD5) concentrations

mimicking the raw water quality of polluted drinking water

sources. This study is unique in the context of NTF research

through its application of a multidimensional investigative

approach. Firstly, nitrification performance was characterized

by measuring traditional bulk physicalechemical indicators

within the influent, effluent and depth-wise filtrate samples

collected throughout NTF bed. Secondly, the microbiology of

biofilm samples collected from within the filter matrix was

characterized and used to link observed changes in biofilm

structure with more traditional bulk chemical performance

parameters. Biofilm samples were characterized through

a number of means. The mass of biofilm protein and carbohy-

drate were closely monitored in order to provide insights into

changes in biopolymer production in response to variations in

organic carbon loading. Direct analytical tools (fluorescence in

situ hybridisation (FISH) and scanning electron microscopy

(SEM)) were also applied to probe the spatial distribution of

ammonia oxidizing bacteria (AOB), nitrite oxidizing bacteria

(NOB) andheterotrophswithin theNTF biofilm. The abundance

of heterotrophs was also monitored using the more traditional

heterotrophic plate count (HPC) method. Finally, NTF perfor-

mance data from this study was also compared with the

performance predicted by the Pearce and Edwards (2010)

empirical design model derived from wastewater applications

under high BOD5 and ammonia loadings in order to assess the

applicability of this design model to potable water NTFs.

2. Materials and methods

2.1. Pilot-plant description

A high flow rate, pilot-scale NTF was constructed at Hope

Valley WTP in South Australia (34�5005400S 138�4101200E) (Fig. 1).

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A more comprehensive description of this NTF has been

published elsewhere (van den Akker et al., 2008). The NTF was

4 m high, 1.5 m in diameter, and supported a 3.0 m bed of TKP

312 polypropylene fill media (2H Plastics, Victoria Australia)

with an area to volume ratio of 240 m�2 m�3. The influent

hydraulic flow rate was maintained at 8 m3 h�1, resulting in

a high hydraulic surface load (per m2 of filter cross-sectional

area) of 125 m3 m�2 d�1. This high hydraulic load was

needed to obtain comparable ammonia and organic mass

loads (g m�2 d�1) to that of conventional wastewater NTFs

(Table 1). Filtrate and biofilm sample ports were positioned

down the vertical axis of the NTF at bed depths of 0.3, 1.5 and

2.7 m from the surface to permit in situ monitoring of the

water chemistry (NHþ4 �N, NO�

2 �N, NO�3 �N, sBOD5, total

organic carbon (TOC)) and biofilm analysis (FISH, SEM, HPC

and total carbohydrate and protein).

2.2. Water quality make-up

The pilot plant was operated under varying ammonia and

organic carbon concentrations to mimic conditions found in

many polluted South East Asian raw water supplies. Varia-

tions in ammonia-N and organic loads were achieved by

dosing ammonium sulphate (Pivot, Australia) and a sucrose

solution (1.5e5% w/v) into the influent, upstream of the NTF’s

rotating distribution arm. A summary of the final influent

ammonia-N and sBOD5 concentrations and corresponding

loading rates are displayed in Table 1.

2.3. Hydraulic tracer analysis

Hydraulic tracer analysis was performed using a fluorescent

dye (rhodamine WT) to investigate the impact that carbon

loading and the subsequent changes in biofilm properties

have on the hydraulic characteristics of the NTF. Tracer

analysiswas performed as described in earlier research by van

den Akker et al. (2008) under both low and high feed sBOD5

concentrations of 1.3 and 8.8 mg L�1. A third recovery

hydraulic tracer study was performed six days following the

termination of carbon (sucrose) dosing. For all experiments,

the hydraulic flow was maintained at 8 m3 h�1. The hydraulic

retention time (HRT) of the NTF was then calculated from the

Fig. 1 e (a) Pilot-scale nitrifying trickling filter; (b) Rotating distribution arm; and (c) Polypropylene media. Arrows point

towards biofilm sample sites positioned at bed depths of 0.3, 1.5 and 2.7 m from the surface.

Table 1 e NTF operating conditions.

NTF NH4eN load sBOD5 load Wetting ratea

(m3 m�2 d�1)Temperature

(�C)Carbon source

g m�2 d�1 mg L�1 g m�2 d�1 mg L�1

Current NTF 0.1e0.90 0.5e5.5 0.22e2.1 1.3e12.1 125 11e22 Sucrose

Pearce and Edwards

(2010)

0.3e0.97 10e31 0.65e1.86 12e43 3.3e84 8e19 Wastewater

(indigenous)

a Flow (m3) m�2 filter cross-sectional plan area d�1.

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recorded tracer timeeconcentration data as described by

Levenspiel (1999).

2.4. Water sampling and chemical analysis

Influent, effluent and depth profile grab sampling (0.5 L) was

performed, on average two to three times per week, with

samples taken at 10:00 am (�2 h). Influent samples were taken

from directly underneath the NTF’s rotating distribution arm.

Filtrate grab samples were taken down the vertical axis of the

NTF bed by inserting a 2.5 cm diameter polypropylene

collection trough horizontally through the each of the biofilm

sample ports. Depth profile samples were collected sequen-

tially from bottom to top within a short 5 min period. Samples

were stored on ice during transport, andwere analysed within

3 h of collection for: ammoniacal-N; nitrite-N; nitrate-N; TOC

and sBOD5 using procedures described previously by van den

Akker et al. (2008). Influent and effluent DO was measured

online using a dissolved oxygen sensor (Danfoss Evita Oxy

Sensor).

2.5. Scanning electron microscopy and fluorescence insitu hybridisation

SEM and FISH were conducted once steady-state nitrification

performance was attained under the respective operating

conditions using the methods described by van den Akker

et al. (2008). For FISH, we used widely published oligonucle-

otide probes specific for Beta-proteobacterial AOB (Nso1225,

50-CGC CAT TGT ATT ACG TGT GA-30), Nitrobacter. (Nit3, 50-CCT GTG CTC CAT GCT CCG-30; Comp NIT3, 50-CCT GTG CTC

CAG GCT CCG-30), Nitrospira (Ntspa662, 50-GGA ATT CCG CGC

TCC TCT-30; Comp Ntspa662 50-GGA ATT CCG CTC TCC TCT-

30) and members of the domain Bacteria (Eub338, 50-GCT GCC

TCC CGT AGG AGT-30). Probes were synthesised and fluo-

rescently labelled with Cy3 at the 50-end by Geneworks

Australia. NON Eub338 (50-GCT GCC TCC CGT AGG AGT-30)was used as a negative control in all FISH experiments. Probe

hybridisation and wash stringencies were obtained from

ProbeBase: http://www.microbialecology.de/probebase/ (Loy

et al., 2003). A stringent wash step followed which involved

counterstaining slides with 2 mg mL�1 of 40, 6-diamidino-2-

phenylindole (DAPI) in accordance with Abell and Bowman

(2005).

2.6. Total protein and carbohydrate

2.6.1. Biofilm sampling and extractionThe distribution of biomass throughout the NTF was charac-

terised by quantifying total protein and carbohydrate per cm2

of media surface area. Media surface samples (1 cm2) were

taken with cotton swabs from each of the three biofilm

samples sites in triplicate, and transported back to the labo-

ratory in 1 mL of distilled water. Biomass was extracted from

the cotton swabs by sonication (10min, 700W, 35 kHz; Cooper

VisionModel 895, Irvine, CA, USA) followed by vigorous vortex

mixing for 2 min. Samples were centrifuged (Phoenix Orbital,

Model 100) at 5000 rpm for 10 min, and the supernatant was

decanted. The extracted biomass pellet was quantified for

total carbohydrate and protein.

2.6.2. AnalysisThe total protein content of the extracted biomass pellet was

determined using the Bradford method (Bradford, 1976).

Extracted biomass was resuspended in 0.1mL of 0.2 M sodium

hydroxide (total volume 0.1 mL) and incubated at room

temperature for 1 h for cell lysis. Bradford protein reagent

(100 mg L�1 coomassie brilliant blue G-250, 5% v/v ethanol,

10% v/v phosphoric acid; 5 mL) was added to the samples and

vortexed. Absorption at 595 nm was measured after 20 min

(Shimadzu, model UV-1700 spectrophotometer) and protein

quantified using bovine serum albumin as standard. The total

carbohydrate content of the extracted biomass pellet was

determined using the phenol reaction method (Hanson and

Phillips, 1981). Extracted biomass was resuspended in 1.0 mL

of distilled water. 1.0 mL of phenol reagent (5.0 g of reagent

grade phenol, 100 mL of distilled water) was added to the

samples and vortexed. Samples were then digested in 5 mL of

reagent grade concentrated sulphuric acid, vortexed and

incubated at room temperature for 15 min. Absorption at

488 nm was measured (HACH, model 2000 spectrophotom-

eter) and total carbohydrate quantified using glucose as

standard. Results are presented as total mass of carbohydrate

or protein per cm2 of filter media surface area.

2.7. Heterotrophic plate count

Heterotrophic bacteria were enumerated from each of the

three biofilms samples sites using the APHA (1992) spread

plate method 9215A. NTF media surface samples (1 cm2) were

taken with sterile cotton swabs in triplicate, and transported

back to the laboratory in 1.0 mL of sterile peptone dilution

water (0.1% peptone, Oxoid Ltd) for immediate analysis.

Aqueous suspension of biomass were extracted from the

swabs by vortex mixing for 5 min, and serially diluted to 10�2

and 10�3 in sterile peptone water. 100 ml aliquots of the two

dilutions water plated in triplicate on R2A agar plates and

incubated aerobically at 30 �C for 48 h. Results were

enumerated by visually counting individual colonies and are

reported in colony forming units (CFU) per cm2 of media

surface area.

2.8. Evaluation of the Pearce and Edwards (2010)nitrification model

This study evaluated the nitrificationmodel develop by Pearce

and Edwards (2010), which is used for the design and opera-

tion of cross-flow plastic media NTFs (Equation (1)). This

model predicts effluent ammonia-N concentrations in

response to hydraulic loading, temperature, ammonia-N and

BOD5 loading. The goal was to determine whether this model

can be applied to the current potable water NTF, which

operated under influent ammonia concentrations some 10-

fold lower and hydraulic loads that were 2e30 times greater

than the wastewater NTFs studied by Pearce and Edwards (see

Table 1). The Pearce and Edwards model was derived from

operational data arising from a number of filter configura-

tions. One of which was not dissimilar to the pilot plant used

in this study, being a 4.5 m deep filter, which operated with

high recirculation rates giving rise to the higher wetting rates

and lower ammonia-N and BOD5 concentrations.

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NH4eNEffluent ¼ 20:81� BOD5 load1:03 �NH4eN load1:52 � Iv�0:36

� T�0:12

(1)

Where:

BOD5 load ¼ g BOD5 m�2 media surface area per day

NH4eN load ¼ g NH4eN m�2 media surface area per day

Iv ¼ media irrigation rate specific to media surface area as

L m�2 media surface area per day

T ¼ Filter effluent temperature (�C)

3. Results and discussion

3.1. Effects of organic carbon on NTF performance

Fig. 2a shows the impact of sucrose induced sBOD5 loads on

effluent ammonia concentrations, for periods when influent

ammonia was maintained at 0.5, 2.0 and 5.0 mg N L�1. Results

show that small increase in sBOD5 adversely impacted on the

nitrification capacity of the NTF, where organic loads greater

than 0.9 g sBOD5 m�2 d�1 caused a significant breakthrough of

ammonia into the filters effluent. Fig. 2b and c show the

impact of sBOD5 loading on nitrification and heterotrophic

activity measured down the NTF bed. Nitrification and carbon

removal rates were calculated for bed depths 0.3, 1.5, 2.7 and

3.0 m by measuring the difference in ammonia-N and TOC

concentrations between these adjacent sample depths and

were calculated as a function of media surface area. Influent

ammonia-Nwasmaintained at 5.0 (�0.2)mg L�1. At the lowest

applied sBOD5 load, a rapid decline in the nitrification rate

down the filter bed can be seen to occur as the process

becomes increasingly ammonia-limited. The introduction of

small amounts of sBOD5 decreases these rates significantly,

resulting in a closing up of the nitrification rates through the

filter depth. TOC removal rates measured down the bed

increased significantly in response to sBOD5 loading e even at

the lowest bed segment of 2.7e3.0 m. This segment however,

was the only area of the filter where nitrification rates had

remained unchanged; most likely because nitrification (at this

depth) was ammonia-limited to begin with.

3.2. Pearce and Edwards (2010) nitrification model

Performance data presented in Fig. 2a are compared side-by-

side with predicted values from the Pearce and Edwards

(2010) nitrification model, which was derived from munic-

ipal wastewater NTFs supplied with higher strength water

(mg L�1 of BOD5 and NH4eN) and operated under lower

hydraulic loading rates than the NTF employed in this study

(Table 1). This model was also based on wastewater BOD5

which is a far more complex matrix than the sucrose solution

used in this study and likely to comprise of numerous

elements with varying rates of biodegradability. Despite these

marked differences, the model accurately predicted the

nitrification performance of the NTF. The sensitivity of nitri-

fication to small changes in sBOD5, as well as the transition

from zero order to first order kinetics for ammonia removal

(e.g. Fig. 2b), underlines the difficulty of accurately predicting

nitrification; hence the requirement for accurate mechanistic

modelling of large amounts of data to allow model fitting and

calibration. Conversely, the simple empirical Pearce and

Edwards (2010) model used in this study predicted the filter

performance well without the need for calibration.

3.3. Heterotrophic plate count and biopolymers

3.3.1. Effects of organic loadFig. 3 shows the impact of organic carbon loading (mg

sBOD5 L�1) on biofilm carbohydrate, protein and HPC,

measured at NTF bed depths of 0.3, 1.5 and 2.7 m. Of these

three parameters, the mass of biofilm carbohydrate respon-

ded the most to carbon loading. A linear relationship between

biofilm carbohydrate and influent sBOD5 concentration was

evident at all biofilm sample sites of 0.3, 1.5 and 2.7 m

(respective Spearman’s rank coefficients (rs) ¼ 0.82, 0.76, 0.80;

p < 0.01). Conversely, the mass of total protein remained

stable, despite the ten-fold increase in carbon substrate

(respective rs ¼ 0.22, 0.37, 0.31; p > 0.05). The stable biofilm

protein and increase in carbohydrate suggest that the accu-

mulation of carbohydrate was extracellular and that the

microbial utilisation of carbon was orientated towards the

luxurious production of polysaccharides, rather than cell

Fig. 2 e (a) Relationship between organic loading and effluent ammonia-N; (b) The impact of sBOD5 on the nitrification rate

measured through the NTF bed; and (c) Relationship between influent TOC and carbon removal rate.

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growth. Examination of the biofilm using SEM confirmed the

development of a thick extracellular polysaccharide ‘crust’

(see Fig. 1 in Supplementary Information).

The increase in biofilm carbohydrate is a characteristic

sign that carbon was in excess and/or that nitrogen and/or

phosphorous were limited. Similar observations have been

made in suspended growth, biological nutrient removal (BNR)

reactors treating domestic and industrial wastewaters with

high concentrations of available carbon (Stover, 1980; Bura

et al., 1998; Durmaz and Sanin, 2001, 2003; Alves et al., 2002,

van den Akker et al., 2010). In the current study, it was more

likely that the biofilm was deficient in phosphorous rather

than nitrogen. This was simply because the concentration of

total phosphorous in the feed water was very low

(<0.04 mg L�1), which resulted in influent sBOD5/P and TOC/P

ratios in excess of 200:1 and 500:1 respectively. Our results

also showed that when the NTF was operated under fixed

influent sBOD5 concentrations of 5.0 mg L�1, a increase in

influent ammonia concentration from 0.5 to 5.0 mg N L�1

(effectively decreasing the C/N ratio from 10 to 1) did not

significantly decrease themass of biofilm carbohydrate (t-test,

p > 0.05), which further supports the hypothesis that the

biofilm was phosphorous-limited.

A small, yet significant relationship between HPC and

influent sBOD5 was evident only at the upper bed depths of 0.3

and 1.5 m, which were areas of the filter that were exposed to

the highest concentrations of sBOD5 (respective rs ¼ 0.52, 0.56;

p < 0.05). At these depths, heterotrophs increased by a factor

of only ca 0.5 log10 units, despite the ten-fold increase in

carbon substrate. The absence of this relationship at 2.7 m

(rs ¼ 0.48; p > 0.05) was surprising given the large increase in

the TOC removal rate that was seen at this depth (see Fig. 2c).

3.3.2. Biopolymers and HPC versus nitrification performanceThe large increase in total biofilm carbohydrate (measured

at 0.3, 1.5 and 2.7 m) also correlated well with a decline in

the overall nitrification capacity (measured as percentage

ammonia-N removal) (respective rs ¼ �0.72, �0.61, �0.62;

p < 0.01). Similar relationships between ammonia removal

and HPC or protein were not evident ( p > 0.05), and therefore

provide circumstantial evidence to suggest that the highmass

of carbohydrate (which implies high polysaccharide produc-

tion) may have be the dominant causative factor for the

decline in nitrification. The reason for this may be related to

the ability of exopolysaccharides to strongly influence the

mass transfer properties of biofilms (Characklis and Marshall,

1990 pp. 118). The high mass of biofilm polysaccharide may

have caused mass transfer limitations through the biofilm,

which subsequently impeded the filter’s ability to maintain

nitrification. The negative relationship between carbohydrate

and nitrification was a unique finding, which has not been

reported for traditional NTFs treating municipal wastewater.

3.3.3. Temporal changes in biofilm carbohydrate andnitrificationFig. 4 shows temporal changes in total biofilm carbohydrate,

effluent DO and nitrification performance (% ammonia-N

removal) in response to a sudden increase in carbon loading.

For this experiment, the carbon load was increased well

beyond native concentrations of 1.5 (�1.0) to 8.8 (�2.9) mg

sBOD5 L�1 (1.5 � 0.5 g sBOD5 m

�2 d�1) for 12 consecutive days

via the addition of sucrose. Influent ammoniawasmaintained

at 5.0 (�0.2) mg N L�1. Results show a steady increase in the

mass of total carbohydrate within the upper bed depths of 0.3

and 1.5 m, within 12 d of increasing the organic carbon load.

The mass of carbohydrate measured at the lowest bed depth

of 2.7 m remained relatively stable due to the high removal of

carbon substrate in the upper depths (Fig. 2c). An inverse

relationship between ammonia removal (%) and the mass of

biofilm carbohydrate was clearly evident, whereby an

increase in the mass of biofilm carbohydrate coincided with

a ten-fold reduction in the NTF’s nitrifying capacity. Once the

sucrose dosing was terminated (day 12), the mass of carbo-

hydrate returned to original steady-state levels within 8e10

days, and this coincided with the complete recovery in nitri-

fication. The rapid decline in biofilm carbohydrate can be

Fig. 3 e Relationship between influent sBOD5 concentration and biofilm carbohydrate, protein and HPC. All data points are

mean of 3 samples ± 1SD. Broken lines represent 95% confidence intervals.

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attributed to degradation by the heterotrophic community,

since carbohydrates have been shown to serve as a usable

source of organic carbon in BNR reactors (Obayashi and

Gaudy, 1973; Stover, 1980; Zhang and Bishop, 2003). Results

also show that effluent DO concentrations responded well to

carbon loading. Once carbon dosing had commenced, effluent

DO progressively decreased from 9.5 to 5.5mg L�1 within 9e10

days. The de-oxygenation of the effluent in response to carbon

loading is characteristic of heterotrophic activity and may

confirm the substantial development of heterotrophs, which

were otherwise underestimated by the HPC technique (see

Fig. 3). This observation highlights the need to use direct

analytical tools such as FISH and SEM to accurately determine

the extent of heterotroph abundance.

3.4. In situ analysis of the biofilm using FISH and SEM

FISH analysis showed that the oligonucleotide probe Nso1225

was successful in identifying the beta sub-group of AOB Pro-

teobacteria, which included species from the genera Nitro-

somonas andNitrosospira. NOB identified in this study belonged

to the genera Nitrospira and also the less common Nitrobacter,

which were identified using oligonucleotide probes Ntspa662

and Nit3 respectively. When the NTF was operated under low

carbon loads (<0.5 g sBOD5 m�2 d�1) the biofilm was thin and

densely packed with large colonies of nitrifiers (see Fig. 2 in

Supplementary Information). Microscopic examination

showed that the biofilm thickness at 0.3 m was 496 (�109) mm

(n ¼ 12 samples). Due to a thin coverage of biomass at the

lower filter depths (1.5 and 2.7 m), biofilm thickness

measurements were difficult to obtain. Biofilms appeared to

be homogenous in structure, dominated by many large

bacterial colonies of AOB and NOB that were distributed

throughout the sub- and peripheral-depths of the biofilm.

Nitrifiers were abundant at all filter bed depths, representing

the majority of DAPI positive microorganisms.

In situ examination of the biofilm using FISH and SEM

showed that a small increase in influent sBOD5 from 1.5 to

5e8 mg L�1 not only increased the biofilm carbohydrate

content, but also promoted a rapid change in the population

structure (see Fig. 3 in Supplementary Information). Within

6e8 days of operating under high carbon loads, the biofilm

thickness increased, measuring 986 (�106), 823 (�63), and 657

(�95) mm at bed depths of 0.3, 1.5, 2.7 m respectively (n ¼ 3

samples). SEM and FISH revealed a more complex heteroge-

nous biofilm structure. Depth-wise stratification of nitrifiers

and heterotrophs was observed at all filter bed depths, which

coincided with the impediment of nitrification throughout

most of filter column. Fewer AOB and even less NOB were

identified. Nitrifiers that were detected were found restricted

deep within the biofilm at depths of 100e980 mm below the

biofilm surface. Studies which have profiled oxygen within

nitrifying biofilms using microelectrodes have found these

depths to be oxygen limited, which can explain the decrease

in nitrification seen here (Schramm et al., 1996; Okabe et al.,

1999, 2002). The outer heterotrophic layer was 93e423 mm

thick and was comprised mostly of filamentous bacteria. The

positive signal obtained from the Eub338 probe, which is

specific for most bacteria, confirmed that the outer filamen-

tous mat was in fact bacterial, not fungal mycelium. These

findings confirmed that the culture dependant HPC method

underestimated the development of heterotrophs, simply

because the HPC technique is limited to bacteria that are able

to form colonies. The high abundance of filamentous bacteria

may have been promoted by the carbon source used in this

study (sucrose), since it has been reported that energy-rich,

sugar based organic substrates can promote filamentous

growth in suspended growth reactors (Chudoba et al., 1985;

Moy et al., 2002).

Observations presented here through the application of

FISH and SEM methods have enabled us to specifically link

biofilm structure with NTF functionality, thereby physically

Fig. 4 e Temporal changes in ammonia removal performance (%), effluent DO and biofilm carbohydrate (cmL2), in response

to a sudden increase in organic load. Data points represent parameter means ± 1 SD of 3 samples.

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verifying what has previously been inferred from many

years of empirical modelling efforts on NTFs (e.g. Parker

and Richards, 1986; Pearce and Williams, 1999; Pearce

and Edwards, 2010) and less comprehensive experiments.

Furthermore, two decades of mathematical modelling work

aimed at understanding microbial competition in mixed

population biofilms has provided a theoretical basis for the

phenomena we have demonstrated here (e.g. Wanner and

Gujer, 1986; Rittmann and Manem, 1992; Gujer, 2010). Hence,

this work effectively bridges the gap between the outcomes of

empirically-based research and the observations inferred

from macroscale physiochemical studies.

3.5. Hydraulic tracer analysis

Hydraulic tracer experiments were performed to characterize

the patterns of flow and to compare the actual HRTs of the NTF

when operated under low and high organic loads (Fig. 5). The

residence time distribution (RTD) curve obtained under a low

organic load was characterised by a short delay in the arrival of

the maximum asymmetric fluorescence peak, followed by

a relatively rapid but steady fluorescence decline within

a pronounced tail. This revealed that flow patterns were repre-

sentative of well dispersed reactor hydraulics. Conversely, the

RTD curve obtained under high organic loads was more drawn-

out and has a longer tail, which resulted in a 40% increase in the

HRT (from 3.8 to 6.4 min). For all tracer experiments, the

hydraulic flow rate was kept constant, therefore the increase in

HRT can be attributed to a thicker biofilm (e.g. Riemer et al., 1980;

Stevens et al., 1986). It was also likely that the increase in biofilm

carbohydrate contributed to the increase in HRT, since extra-

cellular polysaccharides has the ability to retain water (Chenu

and Roberson, 1996). These results suggest that changes in

filter RTD curves and HRT can be a useful parameter for moni-

toring general filter ‘health’ and could also indicate when

preventativemeasures should be taken to avoid filter clogging or

even collapse. One measure to maintain favourable hydraulic

characteristics may include increasing the hydraulic loading

which can induce biomass sloughing (Howell and Atkinson,

1976). A tracer experiment which was performed six days after

the termination of carbon (sucrose) dosing, showed that the

filter had recovered and regained normal hydraulic character-

isticswithout the need to induce sloughing (HRT¼ 3.7min). This

observation coincided with a decline in biofilm carbohydrate

and complete recovery in nitrification throughout the entire

filter bed.

4. Conclusions

This study presents a comprehensive and coherent set of

experimental results on the structure and function of biofilms

in a high-rate NTF designed for potable water treatment. Both

traditional and advance analytical tools were used to link

biofilm composition and structure with NTF performance.

Specific conclusions from this study were:

� Increases in sBOD5 loading adversely impacted on the

nitrification capacity of the NTF. The decline in nitrification

was remarkably similar to predicted model outputs derived

from wastewater NTFs;

� The application of FISH and SEM methods confirmed that

increases in sBOD5 promoted depth-wise stratification of

nitrifiers and heterotrophs within biofilms, thereby physi-

cally validating previous empirical modelling hypotheses;

� The use of a culture dependant spread plate technique

underestimated the development of heterotrophic bacteria

within the biofilm;

� Small increase in influent BOD5 caused a rapid increase in

biofilm carbohydrate, which was an indication of luxurious

exopolysaccharide production in response to nutrient limi-

tation. This was a unique finding that has not been reported

for trickling filters treating wastewater;

� The increase in biofilm carbohydrate correlated well with

the decline in nitrification. It was possible that the high

polysaccharide content of the biofilm caused diffusional

resistance in addition to that caused by heterotrophs; and

� Changes in biofilm characteristics affected the HRT of the

NTF.

Acknowledgements

This research project was funded by Thames Water (gs1) and

supported by United Water International as part of their

commitment to innovation and responsible water

management. The authors would like to thank Dr Guy Abell

and the staff form Hope Valley WTP for their technical

expertise, Dr Michael Short for assisting with pilot plant

construction and Kerry Gascoigne and Michelle Norman for

assisting with biofilm preparation and image analysis.

Appendix. Supplementary data

Supplementary data related to this article can be found online

at doi:10.1016/j.watres.2011.04.017.

Fig. 5 e Tracer analysis comparing residence time

distribution curves recorded under low and high organic

loads.

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r e f e r e n c e s

Abell, G.C.J., Bowman, J.P., 2005. Ecological and biogeographicrelationships of class Flavobacteria in the Southern Ocean.FEMS Microbiology Ecology 51 (2), 265e277.

Ahmad, R., 2005. Synthetic and Natural Organic Removal byBiological Filtration. John Wiley & Sons, Inc.

Alves, C.F., Melo, L.F., Vieira, M.J., 2002. Influence of mediumcomposition on the characteristics of a denitrifying biofilmformed by Alcaligenes denitrificans in a fluidised bed reactor.Process Biochemistry 37 (8), 837e845.

APHA, 1992. Standard Methods for the Examination of Water andWastewater, eightheenth ed. American Public HealthAssociation/American Waterworks Association/WaterEnvironment Federation, Washington, DC, USA.

Biesterfeld, S., Russell, P., Figueroa, L., 2003. Linking nitrifyingbiofilm structure and function through fluorescent in situhybridization and evaluation of nitrification capacity. WaterEnvironment Research 75 (3), 205e215.

Boller, M., Gujer, W., 1986. Nitrification in tertiary trickling filtersfollowed by deep bed filters. Water Research 20 (11),1363e1373.

Bradford, M.M., 1976. A rapid and sensitive method for thequantitation of microgram quantities of protein utilizing theprinciple of protein-dye binding. Analytical Biochemistry 72(1e2), 248e254.

Bura, R., Cheung, M., Liao, B., Finlayson, J., Lee, B.C., Droppo, I.G., Leppard, G.G., Liss, S.N., 1998. Composition ofextracellular polymeric substances in the activated sludgefloc matrix. Water Science and Technology 37 (4e5),325e333.

Characklis, W.G., Marshall, K.C., 1990. In: Characklis, William G.,Kevin, C. (Eds.), Biofilms. Wiley, New York.

Chenu, C., Roberson, E.B., 1996. Diffusion of glucose in microbialextracellular polysaccharide as affected by water potential.Soil Biology and Biochemistry 28 (7), 877e884.

Chudoba, J., 1985. Control of activated sludge filamentousbulking. VI. Formulation of basic principles. Water Research19 (8), 1017e1022.

Datta, T., Racz, L., Kotay, S.M., Goel, R., 2011. Seasonal variations ofnitrifying community in trickling filter-solids contact (TF/SC)activated sludge systems. Bioresource Technology 102 (3),2272e2279.

De Vet, W.W.J.M., Rietveld, L.C., Van Loosdrecht, M.C.M., 2009.Influence of iron on nitrification in full-scale drinking watertrickling filters. Journal of Water Supply: Research andTechnology e AQUA 58 (4), 247e256.

Durmaz, B., Sanin, F.D., 2001. Effect of carbon to nitrogen ratio onthe composition of microbial extracellular polymers inactivated sludge. Water Science and Technology 44 (10),221e229.

Durmaz, B., Sanin, F.D., 2003. Effect of carbon to nitrogen ratio onthe physical and chemical properties of activated sludge.Environmental Technology 24 (11), 1331e1340.

Gujer, W., 2010. Nitrification and me e a subjective review. WaterResearch 44 (1), 1e19.

Hanson, R., Phillips, J., 1981. Chemical composition. In: Manual ofMethods for General Bacteriology by Philipp BacteriologyeLaboratory Manuals. American Society for Microbiology,Washington D.C.

Howell, J.A., Atkinson, B., 1976. Sloughing of microbial film intrickling filters. Water Research 10 (4), 307e315.

Levenspiel, O., 1999. Chemical Reaction Engineering, third ed.John Wiley and Sons Inc., USA.

Loy, A., Horn, M., Wagner, M., 2003. ProbeBase: an online resourcefor rRNA-targeted oligonucleotide probes. Nucleic AcidsResearch 31 (1), 514e516.

Lydmark, P., Lind, M., Sorensson, F., Hermansson, M., 2006.Vertical distribution of nitrifying populations in bacterialbiofilms from a full-scale nitrifying trickling filter.Environmental Microbiology 8 (11), 2036e2049.

Moy, B.Y.P., Tay, J.H., Toh, S.K., Liu, Y., Tay, S.T.L., 2002. Highorganic loading influences the physical characteristics ofaerobic sludge granules. Letters in Applied Microbiology 34 (6),407e412.

Obayashi, A.W., Gaudy Jr., A.F., 1973. Aerobic digestion ofextracellular microbial polysaccharides. Journal (WaterPollution Control Federation) 45 (7), 1584e1594.

Okabe, S., Naitoh, H., Satoh, H., Watanabe, Y., 2002. Structure andfunction of nitrifying biofilms as determined by moleculartechniques and the use of microelectrodes. Water Science andTechnology 46 (1e2), 233e241.

Okabe, S., Satoh, H., Watanabe, Y., 1999. In situ analysis ofnitrifying biofilms as determined by in situ hybridization andthe use of microelectrodes. Applied and EnvironmentalMicrobiology 65 (7), 3182e3191.

Parker, D.S., Richards, T., 1986. Nitrification in trickling filters.Journal (Water Pollution Control Federation) 58 (9),896e902.

Pearce, P., Edwards, W., 2010. A design model for nitrification onstructured cross flow plastic media trickling filters. Water andEnvironment Journal. doi:10.1111/j.1747e6593.2010.00218.x.

Pearce, P., Williams, S., 1999. A nitrification model for mineral-media trickling filters. Journal of the Chartered Institution ofWater and Environmental Management 13 (2), 84e92.

Persson, F., Wik, T., Sorensson, F., Hermansson, M., 2002.Distribution and activity of ammonia oxidizing bacteria ina large full-scale trickling filter. Water Research 36 (6),1439e1448.

Riemer,M.,HolmKristensen,G.,Harremoes, P., 1980.Residence timedistribution in submergedbiofilters.Water Research 14, 949e958.

Rittmann,B.E.,Huck,P.M., 1989.Biological treatmentofpublicwatersupplies. Critical Reviews in Environmental Control 19 (2).

Rittmann, B.E., Manem, J.A., 1992. Development and experimentalevaluation of a steady-state, multispecies biofilm model.Biotechnology and Bioengineering 39 (9), 914e922.

Rittmann, B.E., Snoeyink, V.L., 1984. Achieving biologically stabledrinking water. Journal of the American Water WorksAssociation 76 (10), 106e114.

Satoh, H., Okabe, S., Norimatsu, N., Watanabe, Y., 2000.Significance of substrate C/N ratio on structure and activity ofnitrifying biofilms determined by in situ hybridization and theuse of microelectrodes. Water Science and Technology 41(4e5), 317e321.

Sayre, I.M., 1988. International standards for drinking water.Journal of the AmericanWaterWorks Association 80 (1), 53e60.

Schramm, A., Larsen, L.H., Revsbech, N.P., Ramsing, N.B.,Amann, R., Schleifer, K.H., 1996. Structure and function ofa nitrifying biofilm as determined by in situ hybridization andthe use of microelectrodes. Applied and EnvironmentalMicrobiology 62 (12), 4641e4647.

Stevens, D.K., Berthouex, P.M., Chapman, T.W., 1986. The effect oftracerdiffusion in biofilm on residence time distribution.Water Research 20, 369e375.

Stover, E.L., 1980. Start-up problems at a plant treating food-processing wastewater. Journal of the Water Pollution ControlFederation 52 (2), 249e256.

Tekerlekopoulou, A.G., Papazafiris, P.G.D., Vayenas, D.V., 2010. Afull-scale trickling filter for the simultaneous removal ofammonium, iron and manganese from potable water.Journal of Chemical Technology and Biotechnology 85 (7),1023e1026.

Tekerlekopoulou, A.G., Vayenas, D.V., 2007. Ammonia, iron andmanganese removal from potable water using trickling filters.Desalination 210 (1e3), 225e235.

wat e r r e s e a r c h 4 5 ( 2 0 1 1 ) 3 4 8 9e3 4 9 8 3497

Author's personal copy

Tekerlekopoulou, A.G., Vayenas, D.V., 2008. Simultaneousbiological removal of ammonia, iron and manganese frompotable water using a trickling filter. Biochemical EngineeringJournal 39 (1), 215e220.

van den Akker, B., Beard, H., Kaeding, U., Giglio, S., Short, M.D.,2010. Exploring the relationship between viscous bulking andammonia-oxidiser abundance in activated sludge:a comparison of conventional and IFAS systems. WaterResearch 44 (9), 2919e2929.

van den Akker, B., Holmes, M., Cromar, N., Fallowfield, H., 2008.Application of high rate nitrifying trickling filters for potablewater treatment. Water Research 42 (17), 4514e4524.

Wanner, O., Gujer, W., 1986. Multispecies biofilm model.Biotechnology and Bioengineering 28 (3), 314e328.

Zhang, T.C., Fu, Y.C., Bishop, P.L., 1994. Competition in biofilms.Water Science and Technology 29 (10e11), 263e270.

Zhang, X., Bishop, P.L., 2003. Biodegradability of biofilmextracellular polymeric substances. Chemosphere 50 (1), 63e69.

wat e r r e s e a r c h 4 5 ( 2 0 1 1 ) 3 4 8 9e3 4 9 83498

Appendix. Supplementary data a b

Fig. 1 – (a) Scanning electron micrograph showing the surface structure of a nitrifying biofilm sampled from the NTF (0.3 m) when operated under high organic loads of 5.0 mg sBOD5 L-1; and (b) a higher magnification of image (a). Both images show microorganisms embedded in a thick exopolysaccharide-like crust.

a b c

Fig 2. – In situ structure of the nitrifying biofilm sampled from a NTF bed depth of 0.3 m when operated under low organic loads of 1.3 (±0.5) mg sBOD5 L-1. (a) Scanning electron micrograph of the nitrifying biofilm showing bacteria located on the outer biofilm surface; (b) corresponding fluorescence micrograph of image (a) showing a biofilm cross-section stained with DAPI (blue); and (c) a higher magnification of image (b), showing AOB after fluorescence in situ hybridisation using probe Nso1225 (red). Dashed line represents the media–biofilm interface.

a b

c d

e f g

Fig 3. – In situ structure of biofilms sampled from the NTF when operated under high influent sBOD5 concentrations within the range of 5–8 mg L-1. (a-b) Biofilm sampled at 0.3 m: (a) Scanning electron micrograph of the biofilm surface structure and (b) corresponding fluorescence micrograph depicting the outer heterotrophic filamentous mat targeted with Eub338 (red); (c-d) Biofilm sampled at 1.5 m: (c) Scanning electron micrograph of the biofilm surface structure and (d) corresponding fluorescence micrograph depicting the outer heterotrophic filamentous mat targeted with Eub338 (red); and (e-g) Biofilm sampled at 2.7 m: (e) biofilm cross-section stained with DAPI (blue), (f) corresponding fluorescence micrograph depicting the outer heterotrophic filamentous mat targeted with Eub338 (red), and (g) biofilm cross-section showing the spatial stratification of Nitrobacter targeted with Nit3 (green) and filamentous heterotrophs stained with DAPI (blue). White arrows point towards the biofilm-bulk fluid interface and the dashed line represents the media–biofilm interface.