Water Research 38 (2004) 1197–1206

ARTICLE IN PRESS

*Correspond

7247-826-858.

E-mail addr

(T. Schwartz).

0043-1354/$ - se

doi:10.1016/j.w

Investigation of natural biofilms formed during the productionof drinking water from surface water embankment filtration

Farahnaz Emtiazi, Thomas Schwartz*, Silke Mareike Marten,Peter Krolla-Sidenstein, Ursula Obst

Department of Environmental Microbiology, Forschungszentrum Karlsruhe GmbH, Institute for Technical Chemistry–Water Technology

and Geotechnology Division, P.O. Box 3640, Karlsruhe D-76021, Germany

Received 5 June 2003; received in revised form 8 October 2003; accepted 13 October 2003

Abstract

Populations of bacteria in biofilms from different sites of a drinking water production system were analysed.

Polymerase chain reaction (PCR) and denaturing gradient gel electrophoresis (DGGE) analyses revealed changing

DNA band patterns, suggesting a population shift during bank filtration and processing at the waterworks. In addition,

common DNA bands that were attributed to ubiquitous bacteria were found. Biofilms even developed directly after

UV disinfection (1–2m distance). Their DNA band patterns only partly agreed with those of the biofilms from

the downstream distribution system. Opportunistic pathogenic bacteria in biofilms were analysed using PCR and

Southern blot hybridisation (SBH). Surface water appeared to have a direct influence on the composition of biofilms

in the drinking water distribution system. In spite of preceding filtration and UV disinfection, opportunistic pathogens

such as atypical mycobacteria and Legionella spp. were found in biofilms of drinking water, and Pseudomonas

aeruginosa was detected sporadically. Enterococci were not found in any biofilm. Bacterial cell counts in the

biofilms from surface water to drinking water dropped significantly, and esterase and alanine-aminopeptidase activity

decreased. b-glucosidase activity was not found in the biofilms. Contrary to the results for planktonic bacteria,

inhibitory effects were not observed in biofilms. This suggested an increased tolerance of biofilm bacteria against toxic

compounds.

r 2003 Elsevier Ltd. All rights reserved.

Keywords: Biofilms; Population shifts; Opportunistic pathogenic bacteria; Enzyme activities; Molecular-biological techniques

1. Introduction

Bank filtration and artificial groundwater enrichment

are frequently employed for the production of drinking

water. When surface water enters the aquifer via an

underground passage due to potential gradients gener-

ated by wells, this is referred to as bank filtration.

During underground passage, a variety of chemical and

biological processes occur, by which compounds are

ing author. Tel.: +49-7247-826-802; fax: +49-

ess: thomas.schwartz@itc-wgt.fzk.de

e front matter r 2003 Elsevier Ltd. All rights reserve

atres.2003.10.056

reduced and eliminated. Research results obtained with

respect to the hydraulic, physico-chemical, and chemi-

cal/biological processes revealed a good and stable long-

term cleaning efficiency of bank filtration. This also

applies to the removal of particles, pathogens, a number

of organic, and most trace substances [1] except for some

individual organic polar compounds which are persis-

tent in a nearly unrestricted manner [2]. Microorganisms

significantly contribute to cleaning during the under-

ground passage by enzymatic degradation or partial

metabolism of water impurities and by physico-chemical

processes, e.g. adsorption. However, specific manipula-

tion and use of these microbial elimination processes

have failed, and knowledge of the processes is too

d.

ARTICLE IN PRESSF. Emtiazi et al. / Water Research 38 (2004) 1197–12061198

incomplete. Still these biological processes during

natural underground passage are suited as environmen-

tally compatible and low-cost stage for sustainable

drinking water processing. This is not only true for

Central Europe with its large river basins, but also for

many Third World and threshold countries that have to

produce drinking water from highly contaminated

surface waters. Yet few data are available for the

comparative description of function and population

changes of adhesive bacteria (biofilms) during the

underground passage and in the downstream drinking

water production facilities.

Among the major ecological units of aquatic systems,

which affect water quality, are biofilms that cover

practically all accessible wet surfaces. Biofilms may be

composed of algae, bacteria, fungi and other eukaryotic

microorganisms, and cover surfaces, e.g. in storage

basins, filter systems, and drinking water distribution

lines. As biofilms represent the predominating biological

form of life in habitats of water and soil, it is urgently

required to improve the understanding of the structure

and function of these biocoenoses. There is a lack of

methods for the detection of adhesive bacteria in

classical drinking water analysis. Cultivation processes

cover a small part of the natural planktonic population

only [3]. The limitation of nutrients and environmental

stress situations may induce physiological and morpho-

logical changes in many aquatic bacteria. A dormant

status was described for pathogenic bacteria, such

as Campylobacter spp., E. coli, and Legionella pneumo-

phila [4,5] which makes their isolation and identi-

fication problematic. Therefore, cultivation-independent

molecular-biological methods targeting nucleic acids are

required in addition to biochemico-metabolic analyses

[5].

Our investigations covered biofilms from surface

water, raw water after bank filtration, processed

drinking water prior to and after UV disinfection as

well as from the downstream municipal distribution

system. As an alternative to classical cultivation

methods, molecular-biological methods were applied

with the DNA as target molecule, which exists in each

cell irrespective of its physiological state. Oligonucleo-

tide probes developed for a number of environmental

bacteria and pathogens allowed to comprehensively

describe bacterial populations or identify pathogens

without a cultivation pre-enrichment. By means of the

polymerase chain reaction (PCR), denaturing gradient

gel electrophoresis (DGGE), and southern blot hybridi-

sation (SBH), population shifts of the bacterial biofilms

were investigated and the occurrence of facultatively

pathogenic bacteria, such as legionellae, mycobacteria,

and enterococci, was studied. Enzyme activities were

measured in biofilm samples to determine the metabolic

performance and physiological states of the bacterial

biocoenoses.

2. Material and methods

2.1. Biofilm sampling method

For biofilm formation in drinking water distribution

systems, special devices were used [6,7]. The core of these

devices consisted of a hollow stainless steel cylindrical

element, where stainless steel bolts holding steel platelets

for biofilm growth were screwed into place. The platelet

(15mm� 30mm� 2.5mm) was attached to the end of a

bolt with a small screw. To study natural biofilms, these

devices were installed at different sampling points in the

waterworks, i.e. downstream of the granular activated

carbon filtration (GAC) and downstream of UV

disinfection (DIS). Two devices were installed in house

branch connections within the municipal drinking water

distribution system (DW1 and DW2) 1–2 km down-

stream of the waterworks. Additionally, platelets

(170mm� 20mm� 2mm) were incubated in surface

water used for embankment filtration (SW, Rhine river)

and raw water (RW) from a well downstream of bank

filtration. To examine the biofilm populations, the

platelets were removed after 3–4 weeks, the biofilms

were scraped from the surface using a sterile cell scraper

(Nunc, Wiesbaden, Germany) and suspended in 10ml

water from the sampling site. The biofilm suspensions

were centrifuged for 10min at 10 000g. The pellet was

resuspended in 1ml sterile water. DNA was extracted

from biofilms from the surface water and horizontal well

(QIAamps DNA Mini Kit-50, Qiagen). These DNA

preparations were used as template in PCR. The biofilm

suspensions from GAC, DIS, and DW were used

without extraction for subsequent molecular-biological

analysis.

2.2. PCR-DGGE analysis of different aquatic biofilms

PCR primers targeting the regions (V1-3) of 16S

rDNA of bacteria were used for biofilm analysis

(Table 1) [8,9]. For DGGE analysis, a sequence of

multiple guanines (G) and cytosines (C) was attached to

the 50end of the forward primer [10]. A ‘‘touch-down’’

PCR profile published by Kilb et al. [9] was applied. The

final 100ml reaction mixture contained 2.5 U of HotStar

Taq-DNA polymerase (Qiagen, Germany), 30 pmol of

each primer, 1�PCR buffer, 1.5mM MgCl2, 200mMdNTPs, and 10 ml biofilm suspension or template DNA.

A GeneAmp PCR System 9700 (Applied Biosystems)

was used for PCR. Aliquots of 5 ml were analysed by

electrophoresis in a 1% agarose gel containing ethidium

bromide to check the sizes and amounts of the

amplicons. PCR products were purified using phenol-

chloroform-isoamyl alcohol (25:24:1 vol.), precipitated

with isopropanol, washed with 70% ethanol, dried, and

resuspended in 20 ml sterile water. DGGE analysis of

PCR products (B526 bp and 349 bp) was performed

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Table 1

PCR primers and SBH probes used

Primer/probes Organism Sequence (50-30) Target site Reference

Primer GC27Fa Bacteria AGAGTTTGATCMTGGCTCAGb 8–27 (16S rDNA) [9]

Primer 517R Bacteria ATTACCGCGGCTGCTGG 534–517 (16S rDNA) [9]

Primer 342R Bacteria CTGCTGCCTCCCGTAG 357–342 (16S rDNA) [8]

Primer-Eub338F Bacteria ACTCCTACGGGAGGCAGC 355–338 (16S rDNA) [8]

Primer-Myc1 Mycobacterium (Genus) AAGGAAGGAAACCCACAC 829–847 (16S rDNA) [12]

Probe-Myc2 Facultative pathogenic mycobacteria CCACCTACCGTCAATC 476–492 (16S rDNA) [12]

Probe-Myc3 Mycobacterium (Genus) TTTCACGAACAACGCGACAA 609–590 (16S rDNA) [12]

Primer-616V Bacteria AGAGTTTGATYMTGGCTCAGb 8–27 (16S rDNA) [6]

Primer-630R Bacteria CAKAAAGGAGGTGATCCb 1542–1529 (16S rDNA) [6]

Probe-Leg-705 Legionella sp. CTGGTGTTCCTTCCGATC 705–722 (16S rDNA) [13]

Probe-Legpneu Legionella pneumophila ATCTGACCGTCCCAGGTT 16S rDNA [14]

Primer-1019V Bacteria TAGCTGGTTCTCYBCGAAb 807–824 (23S rDNA) [15]

Primer-1028R Bacteria CCTTCTCCCGAAGTTACGG 1691–1709 (23S rDNA) [15]

Primer-118V Bacteria TCYGAATGGGGNAAC 121–136 (23S rDNA) [15]

Primer-367R Bacteria CACGTGTYCCGCCGTACTCb 374–394 (23S rDNA) [15]

Probe-DB6 Enterococcus faecium CACACAATCGTAACATCCTA 140–158 (23S rDNA) [15]

Probe-DB8 Enterococcus faecalis TAGGTGTTGTTAGCATTTCG 342–361 (23S rDNA) [15]

Probe-Pae1500 Pseudomonas aeruginosa AATCCGGGGTTTCAAGGC 1500–1517 (23S rDNA) [15]

aGC clamp: 50-CGC CCG CCG CGC CCC GCG CCC GTC CCG CCG CCG CCC CCG CCC C-30.bWobbles according to IUPAC: M means A or C, Y means C or T, K means G or T, B means C, G or T, N means A, C, G or T.

F. Emtiazi et al. / Water Research 38 (2004) 1197–1206 1199

with the D-Code-System (Bio-Rad Laboratories GmbH,

Germany) using polyacrylamide gels containing a

gradient of 40–70% urea for primer combination

GC27F/517R and 50–70% for primer combination

GC27F/342R. 10 ml of the PCR products were loaded

on the gel. DGGE gels were run in 1�TAE buffer

(40mM Tris, 20mM acetate, 1mM EDTA, pH 7.4) at

56�C and 70V for 16 h and stained with SYBRtGreen I

(Sigma). Finally, the stained gels were analysed im-

mediately using the LumiImager Working Stationt

(Roche Diagnostics, Germany). DGGE fingerprints

were scored manually by the presence or absence of

bands, independently of intensity. The number of

bands is supposed to correspond roughly to the number

of the bacterial species in the microbial biofilm.

Community similarities were calculated in a pair-wise

manner using the Dice coefficient: Cs ¼ 2jða þ bÞ�1;where j is the number of bands common to samples A

and B, and a and b are the total numbers of bands in

samples A and B, respectively. This index ranges from 0

(no common bands) to 1 (100% similarity of band

patterns) [11].

Intensively stained bands were excised, gel slices

crushed in 20 ml of sterile water, and equilibrated

overnight at 4�C. 10ml of the DNA extract were re-

amplified by PCR and re-run on DGGE gels to verify

the purity of the PCR re-amplification product. PCR

products were purified with the QIAquick Spin PCR

purification kit (Qiagen, Germany) and sequenced by

GENterprise (Mainz, Germany).

2.3. PCR and southern blot hybridisation (SBH)

All primers and probes used for the identification of

hygienically relevant bacteria in different biofilms are

listed in Table 1. For the identification of mycobacteria

in different biofilms, PCR primer combinations were

used according to Schwartz et al. [12]. The oligonucleo-

tides Myc1 and Myc3 were genus-specific for atypical

mycobacteria, whereas Myc2 was specific for faculta-

tively pathogenic mycobacteria. For the detection of

mycobacteria, semi-specific amplification of 16S rDNA

using the PCR primer Myc1 genus-specific for myco-

bacteria together with the primer Eub338R specific for

most Eubacteria was accomplished. This primer set

amplified 500 bp amplicons. After amplification, PCR

products were hybridised with a specific probe Myc3 for

the detection of atypical mycobacteria and with probe

Myc2 for the detection of facultatively pathogenic

mycobacteria. The stringency of this detection was

adapted for Myc2 with 20% formamide content of the

hybridisation buffer. In modification of the published

PCR protocol, HotStar Polymerase from Qiagen (Hil-

den, Germany) was used.

For identification of Legionella spp., DNA was

amplified using the universal primer combination

616V/630R [6]. The PCR mix was heated up to 95�C

for 15min to activate the polymerase. The profile of the

35 cycles was as follows: Denaturing at 94�C for 1min,

annealing at 50�C for 2min, DNA extension at 72�C for

3min.Probes Leg705 specific for Legionella spp. and

ARTICLE IN PRESSF. Emtiazi et al. / Water Research 38 (2004) 1197–12061200

Legpneu specific for Legionella pneumophila were used

for identification by Southern blot hybridisation [13,14].

To confirm the presence or absence of Legionella spp.

and L. pneumophila, the stringency of the subsequent

hybridisation reaction was evaluated by testing forma-

mide contents ranging from 0%, 15%, 20% to 25% with

cultures of Legionella pneumophila as reference strain.

For high specificity the used formamide concentration

was 15% for probe Leg705 and 25% for probe Legpneu.

Primer combination 1019V/1028R was used for the

amplification of bacterial 23S rDNA prior to the

detection of Pseudomonas aeruginosa. The PCR profile

from Frahm et al. [15] was applied. For the identifica-

tion of Enterococcus faecium and Enterococcus faecalis

bacterial 23S rDNA was first amplified using primer pair

118V and 367R. PCR was carried out as follows:

activation of the Taq polymerase at 95�C for 15min,

then 35 cycles consisting of denaturation at 94�C for

1min, annealing at 44�C for 2min, and extension at

72�C for 3min. For the specific detection of Enter-

ococcus faecium, Enterococcus faecalis, and Pseudomo-

nas aeruginosa, probes DB6, DB8, and Pae1500 were

used, respectively (Table. 1). Different formamide con-

centrations (0%, 20%, 25%, 30%) were tested in the

hybridisation buffer to adjust stringent conditions.

Enterococcus faecium, Enterococcus faecalis and Pseu-

domonas aeruginosa cultures were used as references.

For high specificity the used formamide concentration

was 20% for DB6, 30% for DB8, and 30% for Pae1500.

Southern blot hybridisation was performed according

to Schwartz et al. [12]. Detection was accomplished by a

LumiImager Working Stationt (Roche Diagnostics,

Mannheim, Germany).

2.4. Total bacterial cell count and enzyme activities

Total bacterial cells were counted using DAPI (40,6-

diamidine-20-phenyl-indole-dihydrochloride, Merck,

Germany) staining [6].

Microbial esterase, alanine–aminopeptidase, and b-glucosidase activities and total cell counts were investi-

gated in the different biofilms from all sampling points.

To determine concentration-effect correlations, bio-

films were scraped from the platelet substrates using

sterile cell scrapers (Nunc, Wiesbaden, Germany),

suspended in 10ml PBS (NaCl 8 g, KCl 0.2 g, KH2PO4

0.2 g, and Na2HPO4� 2H2O 1.44 g per litre, pH 7.4)

buffer, and enzyme activities were measured with an

aliquot of 200 ml biofilm suspension in microtiter plates.

The enzyme activity values were calculated as substrate

turnover per hour and cm2 surface area due to the

different sizes of used platelets. Biofilm suspensions were

diluted with sodium chloride solution (0.14mol l�1) in

10% steps from 0 to 100% for dose response experi-

ments. Esterase, alanine–aminopeptidase, and b-glyco-sidase activities were detected using fluorescein diacetate

(final concentration: 2.4� 10�8mol/assay), l-alanin-4-

methoxy-b-naphthylamide (1.2� 10�7mol/assay), and

4-nitrophenyl-b-d-glucopyranoside (3.32� 10�8mol/as-

say) as substrate, respectively [16]. Samples were

incubated at 25�C for 120min for the detection of

esterase and at 30�C for 19 h for the detection of

alanine–aminopeptidase and b-glucosidase. The concen-tration of the released fluorescein and 4-methoxy-b-naphthylamine was determined fluorimetrically at wa-

velengths of 485 nm and 355 nm (Labsystems Fluoros-

kan). The released 4-nitrophenol was measured

photometrically at 405 nm (Labsystems Multiskan MS).

3. Results and discussion

3.1. Population analysis of biofilms by PCR and DGGE

Molecular-biological methods allow to study the

composition of the microflora in natural biofilms of

drinking water conditioning facilities without cultiva-

tion. In comparison to cultivation techniques, charac-

terisation of biofilms by means of PCR and DGGE

analysis is rapid and may be more comprehensive

process. Within the framework of the present study, a

touchdown technique was employed for PCR amplifica-

tion using two different primer combinations. Touch-

down techniques have been considered useful to amplify

also rare DNA targets. Muyzer et al. [8] report that

bacterial populations that make up at least 1% of the

total community can be detected by PCR-DGGE.

In this study, a PCR-DGGE protocol was developed

for the detection of population shifts of bacteria in

different biofilms during the embankment purification

process from surface water to drinking water. The

DGGE band patterns of the PCR product obtained with

the primer set GC27F and 517R are presented in Fig. 1.

The length of the PCR products was 526 bp. After

separating the amplicons by DGGE, the number of

bands varied from 11 to 21, depending on the biofilm

sampling point analysed. DGGE profiles of a 349 bp

PCR product obtained with the primer set GC27F and

342R, were different and showed fewer DNA bands

(data not shown). Bands obtained with the primer set

GC27F/517R were more intense than those with

GC27F/342R. When DGGE profiles generated with

the two primer systems were compared, the similarities

of biofilms decreased again continuously from surface

water to drinking water. In spite of PCR product

purification, a substantial number of weak DNA bands

from less abundant bacteria was generally visible in all

samples independently of the primer set used (Fig. 1,

lanes 2, 7). PCR efficiency may have been influenced by

PCR inhibitors present particularly in the biofilm

samples, resulting in DGGE profiles with weaker

DNA bands and reduced band numbers. It is hence

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Fig. 1. DGGE band pattern of biofilm bacteria from different sampling sites using primer combination GC27F/517R after SYBR

Green staining. Lane 1: Eschericha coli as positive control; lane 2: surface water SW1; lane 3: surface water SW2; lane 4: raw water RW;

lane 5: drinking water after activated carbon filtration GAC; lane 6: drinking water after disinfection DIS1; lane 7: drinking water after

disinfection DIS2; lane 8: distribution system DW1; lane 9: distribution system DW2. Arrows indicate bands excised for sequencing.

F. Emtiazi et al. / Water Research 38 (2004) 1197–1206 1201

pointed out that the different biofilm preparation

techniques (DNA extraction or direct analysis of biofilm

aliquots) may have had an effect on the abundance of

DNA band patterns. Generally, the PCR-DGGE

fingerprints of the biofilms did not exhibit any unique

pattern from one sampling point to the next. Different

patterns were determined even within the distribution

system, although the distribution area was supplied with

the same drinking water (Fig. 1, lanes 8 and 9).

Dice coefficients (Cs) for the pair-wise comparison of

biofilm community composition similarities are listed in

Table 2. As expected, the Cs index for similar aquatic

habitats was higher than the similarity index between

different aquatic compartments, with similarity Cs

values decreasing continuously from surface water to

drinking water. This indicates that surface water

bacteria passing the different conditioning steps also

occur in the subsequent drinking water facilities.

Oligotrophic drinking water systems may be preferred

by certain environmental groups of bacteria from

surface water. The lowest Cs values were measured

between raw water biofilms after embankment filtration

and biofilms within the distribution system. This may be

due to the more anaerobic conditions during under-

ground passage. In some cases, the similarity values

generated with the two primer sets differed considerably.

This may be explained by differences in primer

mismatches between the two PCR systems.

To obtain more detailed information about some of

the community members in these biofilms, a number of

strong DGGE bands (see arrows in Fig. 1) were excised

from the gel, re-amplified, subjected to electrophoresis,

and sequenced. Comparison of the 16S rDNA sequences

with sequences available in GenBank databases indi-

cated that beta-Proteobacteria were most abundant

(similarity between 87% and 99%) (Table 3). Sequences

with high similarity to Dechloromonas (97–99%) were

found in high abundance in the biofilms from raw water,

drinking water after UV disinfection, and the municipal

drinking water distribution system. Sequences with

similarity to Pseudomonas sp. and Acidovorax sp. were

identified in biofilms from surface water only, and

sequences similar to Azospirillum doebereinerae and

Pseudomonas diminuta (96–99%) in biofilms from raw

water and disinfected drinking water. One DNA band of

a gamma-Proteobacterium (99% similarity to Pseudo-

monas marginalis) originated from a biofilm from the

municipal drinking water distribution system. The

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Table 2

Comparison of PCR-DGGE fingerprints from different biofilms by the Dice coefficient

Sample SW RW GAC DIS DW1 DW2

SWa 1.0

SWb 1.0

RWa 0.50 1.0

RWb 0.48 1.0

GACa 0.50 0.29 1.0

GACb 0.46 0.59 1.0

DISa 0.37 0.28 0.47 1.0

DISb 0.33 0.40 0.54 1.0

DW1a 0.32 0.26 0.45 0.46 1.0

DW1b 0.30 0.33 0.33 0.36 1.0

DW2a 0.41 0.37 0.46 0.46 0.56 1.0

DW2b 0.36 0.24 0.38 0.25 0.45 1.0

Two primer combinations GC27F/517R (a) and GC27F/342R (b) were used; for each primer combination 2-3 experiments were

performed per sampling point. SW: surface water; RW: raw water; GAC: downstream of granular activated carbon filters; DIS:

downstream of UV disinfection; DW1 and DW2: within the distribution system.

Table 3

Similarity of sequences of selected DGGE bands, as determined by BLAST nucleotide search

Selected DGGE bandsa Related sequence Similarity (%) Subclass Habitatb

P1 Nitrosomonas oligotropha 97 Beta-Proteobacteria RW

P2 Beta Proteobacterium CRE-PA84 91 Beta-Proteobacteria DW1

P3 Beta Proteobacterium Spb298 99 Beta-Proteobacteria DIS

P4 Pseudomonas marginalis, NZCX27 99 Gamma-Proteobacteria DW2

P5 Azospirillum doebereineri 99 Alpha-Proteobacteria DIS

P6 Sludge bacterium S21 99 Beta-Proteobacteria DIS

P7 Beta Proteobacterium A0640 97 Beta-Proteobacteria RW

P8 Beta Proteobacterium UCT N117 99 Beta-Proteobacteria GAC

P10 Pseudomonas sp. C96E 99 Gamma-Proteobacteria SW

P11 Acidovorax sp. G8B1 97 Beta-Proteobacteria SW

P12 Pseudomonas diminuta 98 Alpha-Proteobacteria RW

P13 Dechloromonas spp. 98 Beta-Proteobacteria DW2

P14 Eubacterium F13.40 94 Beta-Proteobacteria GAC

B1 Bacterium GKS2-174 87 Beta-Proteobacteria DIS

B4b Azospirillum sp. Mat2-1a 95 Alpha-Proteobacteria DIS

B6 Dechloromonas spp. 99 Beta-Proteobacteria DIS

B8 Dechloromonas spp. 98 Beta-Proteobacteria DW2

B9 Bacterium clone IAFDn47 84 Beta-Proteobacteria DW1

B11 Pseudomonas spinosa, ATCC 93 Beta-Proteobacteria SW

B12 Pseudomonas sp. clone Pseud3a 99 Gamma-Proteobacteria SW

B12b Pseudomonas sp. clone Pseud3a 99 Gamma-Proteobacteria SW

B13 Bacterium BVB72 98 Beta-Proteobacteria SW

B15a Dechloromonas spp. 98 Beta-Proteobacteria RW

B17a Dechloromonas spp. 96 Beta-Proteobacteria DW2

B17b Dechloromonas spp. 97 Beta-Proteobacteria DW2

B18a Brevundimonas sp. Dcm7A 99 Alpha-Proteobacteria DW2

B18b Alpha Proteobacterium FL14F11 96 Alpha-Proteobacteria DW2

aBand numbers P1 to P14 were amplified using primer set GC27F/517R (arrows in Fig. 1) and band numbers B1 to B18b were

amplified using primer set GC27F/342R (not shown).bSW, surface water; RW, raw water; GAC, drinking water after activated carbon filtration; DIS, drinking water after disinfection;

DW, drinking water distribution system.

F. Emtiazi et al. / Water Research 38 (2004) 1197–12061202

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results confirm previous studies on biofilms grown at the

same drinking water sampling points within the

distribution system. In situ hybridisation experiments

using subclass-specific labelled probes for Proteobacteria

revealed that beta-Proteobacteria were most frequently

found, but also alpha- and gamma-Proteobacteria could

be detected in significant minor percentages in biofilms

from embankment filtered drinking water [6]. In

drinking water biofilm communities from distribution

systems supplied with conditioned groundwater in

Hamburg, Berlin, Mainz, and drinking water in Stock-

holm, which is gained from surface water, a higher

number of beta-Proteobacteria, particularly Aquabacter-

ium commune, was detected [17]. In our analyses none of

the dominant PCR amplicons showed any significant

homology with A. commune. Kawai et al. [18] reported

constrastingly that alpha- and gamma-Proteobacteria

were dominant in purified water which had been

prepared by ion exchange from tap water.

3.2. Identification of hygienically relevant bacteria in

different biofilms

Bacterial re-growth in biofilms can become a problem

in water distribution systems. In the present study, the

occurrence of hygienically relevant bacteria, such as

facultatively pathogenic mycobacteria, Legionella spp.,

enterococci and Pseudomonas aeruginosa was analysed

by PCR and Southern blot hybridisation (SBH) with

specific primers and probes in biofilms from surface

water to drinking water (Table 1). Saprophytic myco-

bacteria were detected in all samples (Fig. 2b1), and

Fig. 2. Identification of saprophytic and facultatively pathogenic atyp

Eub338R/Myc1; (2b1) SBH with the probe Myc3 for identification of

identification of facultatively pathogenic atypical mycobacteria; lane

water RW; lane 4: drinking water after activated carbon filtration

distribution system DW1; lane 7: distribution system DW2; lanes 8

positive controls.

facultatively pathogenic mycobacteria in the biofilm

from drinking water directly after UV disinfection and

in 1 sample from the distribution system (Fig. 2b2).

Previously, Schwartz et al. [12] demonstrated the

presence of non-tuberculous and non-pathogenic myco-

bacteria in native biofilm samples from the same

drinking water distribution systems. Atypical mycobac-

teria occurred more frequently in biofilms from bank-

filtered drinking water than in biofilms from drinking

water conditioned from groundwater. Hall-Stoodley

et al. [19] found the rapidly growing facultative

pathogens Mycobacterium fortuitum and Mycobacterium

chelonae in soil, freshwater, seawater, wastewater and

potable water. The environment and even drinking

water must hence be considered as a potential source of

infection with facultatively pathogenic mycobacteria.

Bacteria of the genus Legionella and Legionella

pneumophila, the etiologic agent of the Legionnaires’

disease, normally inhabit fresh water or wet soil and can

live as intracellular parasites of amoebae and ciliates or

cyanobacteria. They are found both in natural and man-

made environments, such as cooling towers [20,21]. Lye

et al. [22] detected significant amounts of Legionella spp.

in groundwater and potable water. Legionella spp. and

especially Legionella pneumophila were described to be

adapted to warm water systems, where they multiply

most effectively [20,21]. In larger water distribution

systems Legionella spp. find good survival conditions,

increased temperatures, and nutrients (such as sediments

and biofilms). In the present study, Legionella spp. were

detected using genus-specific probes in biofilms from

surface water to drinking water, but there was no

ical mycobacteria by PCR and SBH: (2a) PCR with primer set

saprophytic mycobacteria; (2b2) SBH with the probe Myc2 for

1: surface water SW1; lane 2: surface water SW2; lane 3: raw

GAC; lane 5: drinking water after disinfection DIS; lane 6:

and 9: Mycobacterium avium and Mycobacterium kansasii as

ARTICLE IN PRESS

Table 4

Enzyme activities and total cell counts in different biofilms

Biofilm samples from Enzyme activities (mol h�1 cm�2) Total cell countsa

Esterase Alanine-aminopeptidase (DAPI counts cm�2)

Surface water 8.8� 10�6 (74.3� 10�7) 4.0� 10�6 (73.4� 10�8) 7.2� 106

Raw water 8.4� 10�6 (71.0� 10�7) 1.9� 10�6 (72.6� 10�7) 5.6� 106

Drinking water after activated carbon filtration 5.7� 10�6 (71.8� 10�7) 3.4� 10�7 (71.7� 10�8) 7.4� 105

Drinking water after UV disinfection 2.7� 10�6 (72.9� 10�7) 2.3� 10�7 (72.0� 10�9) 3.0� 105

Drinking water from the distribution system 4.9� 10�6 (71.1� 10�7) 6.6� 10�7 (72.6� 10�8) 5.7� 105

aStandard error, 20–30%; n ¼ 223:

F. Emtiazi et al. / Water Research 38 (2004) 1197–12061204

evidence for Legionella pneumophila. Using PCR and

SBH techniques, Schwartz et al. [6] also detected

Legionella spp. in different biofilms of a cold drinking

water distribution system.

Pseudomonas aeruginosa is an important pathogen in

nosocomial infections and its frequent presence in

recreational and drinking water is a significant threat

to public health [23]. We detected P. aeruginosa in one

biofilm sample from drinking water, but not in the

whole distribution system. The indicators of fecal

contamination, Enterococcus faecium and Entercoccus

faecalis, were not detected in the investigated biofilms.

In previous molecular biological and conventional

microbiological studies, enterococci were detected only

sporadically after pipe bursts in biofilms from these

drinking water distribution systems [6].

These results indicate that saprophytic as well as

facultatively pathogenic atypical mycobacteria, Legio-

nella spp., and Pseudomonas aeruginosa may have been

transferred from their natural habitat of surface water to

the drinking water distribution system, although water

treatment involved bank filtration and disinfection.

3.3. Microbial cell counts and enzyme activities

Survival and physiology of bacteria depend on

nutrient supply, and this affects the purification

performance during raw water conditioning. Bacterial

counts per biofilm surface area measured by DAPI

staining decreased by 1–2 orders of magnitude from

surface water to raw water and further from non-

disinfected drinking water after filtration to disinfected

drinking water directly after UV disinfection (Table 4)

probably due to filtration and disinfection. In the

downstream water distribution system, bacterial counts

per biofilm surface area were slightly increased.

Enzyme activities of esterases for general metabolic

activities, b-glucosidase for carbohydrate metabolism

and alanine-amino-peptidase for protein metabolism

were measured to physiologically characterise biofilms

in surface water and drinking water treatment steps. In

all biofilm samples, activity of alanine-amino-peptidase

was much lower than of esterase, and glucosidase

activity was very low, indicating a high general cell

and protein but a low carbohydrate metabolism. In

parallel with the DAPI counts, enzymatic substrate

utilisation was decreased after filtration and disinfection

but still detectable (Table 4). For the downstream

distribution system esterase and alanine-amino-pepti-

dase activities slightly increased. The decrease of enzyme

activity was proportional to the decrease of bacterial

numbers. Hence, the biofilms did not differ in specific

enzyme activities.

To identify effects of soluble water components on

biofilm enzyme activities, serial dilutions of the sus-

pended biofilms were tested. There was a linear

correlation between enzyme activities and biofilm

biomass (not shown) indicating non-toxic conditions.

A non-linear relationship was expected, if inhibitors

were present.

Lehman et al. [24] reported increased amino-peptidase

activity of attached organisms compared to unattached

organisms in artificial systems supplied with ground-

water. Enzyme activities depend on bacterial numbers,

water quality, natural substrates, and other environ-

mental factors (e.g. slow sand filtration, bank filtration

of groundwater) [24–26]. Many of these studies focussed

on free-living bacteria without considering attached

biocoenoses. In contrast to previous studies with

planktonic systems, which described dose-dependent

inhibition effects in bulk water [27], no enzyme

inhibition was observed in our biofilm samples. This

suggests that biofilms are more resistant to toxic

compounds than free living bacteria. Miettinen et al.

[25] reported that many xenobiotics inhibited exoenzyme

activities. This inhibition has been used as a biochemical

toxicity test for the pollution of surface waters [16].

Hendel et al. [26] reported that the microbial community

in the sand filter probably acts as a biological buffer

against increases in the loads of organic matter and

nutrients in the recharge plant.

High extracellular enzyme activities reflect high

substrate availability and typically are characteristic of

untreated water. Reductions in cell counts and enzyme

ARTICLE IN PRESSF. Emtiazi et al. / Water Research 38 (2004) 1197–1206 1205

activities of biofilms during the purification process from

surface water to drinking water (Table 4) reflected an

improvement in its trophic status, indicating that the

purification process principally fulfilled its function.

However, the presence of (facultative) pathogens in

biofilms after UV treatment and in the distribution

system is a potential threat to human health. Therefore,

microbiological, molecular biological and physiological

controls are useful tools for monitoring the quality of

drinking water and the efficiency of the conditioning

processes.

4. Conclusions

* The characterisation of biofilms by means of PCR,

DGGE, and sequence analysis is a rapid cultivation-

independent method to gain insight into the micro-

bial population composition of aquatic habitats.* The biofilms of the different drinking water con-

ditioning sampling sites showed different band

patterns, and were apparently made up by different

populations. Even within one compartment, the

biofilms were not unique.* The similarity index in similar aquatic habitats was

higher than the similarity index between different

aquatic compartments, indicating that similar ecolo-

gical niches are inhabited by similar bacterial

populations.* Most of the bacteria identified belonged to beta-

Proteobacteria.* Saprophytic mycobacteria and legionellae were de-

tected in all biofilms studied from surface to drinking

water within the distribution system, indicating a

possible passage of bacteria from surface to drinking

water. Positive Southern Blots for Pseudomonas

aeruginosa and facultatively pathogenic mycobacter-

ia were obtained from some biofilms of drinking

water directly after UV disinfection and from the

downstream distribution system. This potential

source of infection is hygienically of concern,

drinking water may need improvement.* Legionella pneumophila and the indicators of faecal

contamination, Enterococcus faecium and E. faecalis,

were not detected in the biofilms by specific probing.

Data for fecal indicators hence do not allow to draw

conclusions on the presence of pathogenic bacteria in

biofilms.* Effects of toxic compounds, as typically observed for

bulk water samples, were not detected for our biofilm

samples, indicating an increased tolerance of biofilm

communities against toxic compounds.* The reduction in enzyme activities during the

purification process from surface water to drinking

water reflect an improvement in its trophic status.

Acknowledgements

We thank the EU for funding the study (EVK1-CT-

1999-00001). Special thanks are given to Dr. Birgit

Kuhlmann, Dr. Beate Kilb, Dr. Holger Volkmann, and

Erik Ziemann for their helpful support. We also thank

Silke Kirchen for her technical assistance during

sampling and laboratory analyses. We thank the

technical staff of the municipal drinking water supplier

for installing the modified devices.

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