This article was downloaded by:[ANKOS 2007 ORDER Consortium][ANKOS 2007 ORDER Consortium]

On: 3 May 2007Access Details: [subscription number 772814176]Publisher: Taylor & FrancisInforma Ltd Registered in England and Wales Registered Number: 1072954Registered office: Mortimer House, 37-41 Mortimer Street, London W1T 3JH, UK

BiofoulingThe Journal of Bioadhesion and BiofilmResearchPublication details, including instructions for authors and subscription information:http://www.informaworld.com/smpp/title~content=t713454511

A novel application of Fourier-transformed infraredspectroscopy: classification of slime from staphylococci

To cite this Article: , 'A novel application of Fourier-transformed infraredspectroscopy: classification of slime from staphylococci', Biofouling, 23:1, 63 - 71To link to this article: DOI: 10.1080/08927010601143524URL: http://dx.doi.org/10.1080/08927010601143524

PLEASE SCROLL DOWN FOR ARTICLE

Full terms and conditions of use: http://www.informaworld.com/terms-and-conditions-of-access.pdf

This article maybe used for research, teaching and private study purposes. Any substantial or systematic reproduction,re-distribution, re-selling, loan or sub-licensing, systematic supply or distribution in any form to anyone is expresslyforbidden.

The publisher does not give any warranty express or implied or make any representation that the contents will becomplete or accurate or up to date. The accuracy of any instructions, formulae and drug doses should beindependently verified with primary sources. The publisher shall not be liable for any loss, actions, claims, proceedings,demand or costs or damages whatsoever or howsoever caused arising directly or indirectly in connection with orarising out of the use of this material.

© Taylor and Francis 2007

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

A novel application of Fourier-transformed infrared spectroscopy:classification of slime from staphylococci

AYNUR KARADEN_IZL_I1, FET_IYE KOLAYLI1 & KIVANC ERGEN2

1Department of Microbiology, Faculty of Medicine, and 2Department of Biophysics, Faculty of Medicine, Kocaeli University,_Izmit-Kocaeli, Turkey

(Received 11 March 2006; accepted 15 November 2006)

AbstractIt has been proposed that the virulence of nosocomial Staphylococcus infections associated with indwelling medical devices isrelated to the ability of the bacterium to colonise these materials by forming a biofilm composed of multilayered cell clustersembedded in a slime matrix. However, the pathogenic role of exopolysaccharide biofilms is not fully understood. A newmethod was sought for differentiating the structure of slime from two closely related bacterial strains, Staphylococcus aureusand Staphylococcus epidermidis. Using PCR it was confirmed that these strains were positive for the icaA and icaD genes andthe complete ica operon (2.7 kb). Monosaccharide analysis by thin-layer chromatography revealed an identical profile forboth strains, with xylose and glucose present among the four visible bands. Using Fourier-transformed infrared spectroscopyand hierarchical cluster analysis, three of four S. aureus samples (75%), and four of five S. epidermidis samples were groupedaccording to species. A novel FTIR approach in classifying slime produced by S. aureus and S. epidermidis is reported.

Keywords: Slime, Staphylococcus epidermidis, Staphylococcus aureus, FTIR, identification, pathogenicity

Introduction

Staphylococci are generally regarded as opportunis-

tic agents possessing an intrinsic pathogenic char-

acter (Frere et al. 1999; Huebner & Goldmann,

1999). Staphylococcus epidermidis is a saprophyte and

part of the normal mucosa and skin microflora.

Staphylococcus aureus is the causative agent in

numerous invasive and toxigenic diseases. In recent

years, biofilm-forming Coagulase-negative staphy-

lococci (CoNS), particularly S. epidermidis, together

with S. aureus, have emerged as a frequent cause of

nosocomial infections in association with indwelling

medical devices such as intravascular catheters,

cerebrospinal fluid shunts, and prosthetic heart

valves (reviewed in Christensen et al. 1994; Rupp &

Archer, 1994).

For some Staphylococcus strains, the production

of extracellular polysaccharides (EPS), variously

termed ‘‘capsule’’, ‘‘glycocalyx’’ or ‘‘slime sub-

stance’’ is a significant virulence factor (Wilkinson,

1983; Vann et al. 1988; Drewry et al. 1990). The

term biofilm has been defined as a structured com-

munity of bacteria enclosed in slime (MacKintosh

et al. 2006). The virulence of S. epidermidis in

infections presumably reflects its ability to colonise

medical devices by forming a biofilm of multilayered

cell clusters embedded in a slime matrix (Ludwicka

et al. 1984). Slime is mainly polysaccharidic in

nature, and consists of glycosaminoglycans, but also

includes nucleic acids, proteins, minerals, nutrients,

and cell wall material (Costerton et al. 1987; Arciola

et al. 2002). The EPS slime is not a true capsule, but

is loosely bound to the Staphylococus cell (Gotz,

2002). It has been shown that both S. aureus and S.

epidermidis can form slime (Montanaro et al. 1998;

Ammendolia et al. 1999; Arciola et al. 1999;

Crampton et al. 1999; McKenney et al. 1999). In

addition, S. aureus is well known for its ability to

express molecules that recognise host matrix proteins

(Christensen et al. 1982; Barth et al. 1989; Mon-

tanaro et al. 1998; Vaudaux et al. 1990). Thus, there

are two possible explanations for the ability of

Staphylococcus species to colonise artificial materials,

namely, bacterial production of EPS and the

presence of adhesions for matrix proteins that,

in vivo, are adsorbed onto the biomaterial surface

(Christensen et al. 1982; Montanaro et al. 1998;

Correspondence: Kivanc Ergen, Department of Biophysics, Kocaeli University Faculty of Medicine, Umuttepe Yerleskesi, Eski _Istanbul Yolu, 41380

Umuttepe, _Izmit-Kocaeli, Turkey. Fax: þ(90) 262 303 7003. E-mail: ergen68@yahoo.co.uk

Aynur Karadenizli and Fetiye Kolayli contributed equally.

Biofouling, 2007; 23(1): 63 – 71

ISSN 0892-7014 print/ISSN 1029-2454 online � 2007 Taylor & Francis

DOI: 10.1080/08927010601143524

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

Barth et al. 1989). According to these proposed

mechanisms, bacterial adhesion and aggregation,

followed by bacterial proliferation and slime produc-

tion on the implant, results in a biofilm capable of

establishing a virulent and persistent infection

(MacKintosh et al. 2006).

Over the last few years, several studies have sought

to elucidate the structures and pathogenetic mechan-

isms by which staphylococci cause severe and

irreducible infections in association with biomaterials

(Foster & McDevitt, 1994; An & Friedmann, 1998;

Montanaro & Arciola, 2000). Activation of the ica

operon initiates the synthesis of polysaccharide

intercellular adhesion (PIA), which leads to surface

accumulation of cellular aggregates by supporting

cell-to-cell bacterial contact (Arciola et al. 2001,

reviewed in Dunne, 2002). PIA is a surface-

associated, linear b1-6 N-acetyl-D-glucosaminylgly-

can (Feigin et al. 1992). It is a product of the

icaADBC gene cluster and is a virulence factor in the

pathogenesis of foreign-body infections (Hoyle et al.

1993). In the operon, expression of icaA and icaD is

required for full slime synthesis (Gerke et al. 1998).

Fourier-transformed infrared spectroscopy (FTIR)

is a valuable technique in molecular biology due to

its high sensitivity in detecting changes in the func-

tional groups belonging to tissue components such as

lipids, proteins, carbohydrates, and nucleic acids.

The shift in peak positions, change in bandwidths,

and intensity of the bands all provide structural and

functional information of potential diagnostic value

(Severcan et al. 2000; 2005; Toyran et al. 2004).

FTIR has been used for EPS characterisation (Davies

et al. 1993; Allen et al. 2004).

When dealing with infected biomedical implants,

it is important to recognise that organisms in a

biofilm are far more resistant to antimicrobial agents

than are organisms in suspension (Dunne, 2002).

Fuller elucidation of the pathogenic role of slime is

necessary to better understand the infections caused

by slime-producing bacteria. Also of importance is

the ability to differentiate slime characteristics in

drug-resistant biofilms and to identify the strains

from slime scraped off the biomaterial surface

without the need for culture. The present study took

the novel approach of using FTIR spectroscopy to

identify the slime constituents and the differences

between S. aureus and S. epidermidis that are positive

for the icaA and icaD genes and the complete operon

responsible for slime production. These determina-

tions and differentiation could not be made with

other molecular methods such as PCR or analysis

of monosaccharide content. Thin-layer chroma-

tography (TLC) was also used to analyse the

monosaccharide composition of EPS from phenol-

extracted slime preparations (PES) of S. epidermidis

and S. aureus.

Materials and methods

Bacterial isolates

S. epidermidis and S. aureus were clinically isolated

from catheter tips. All isolates were identified by

classical microbiological methods as well as the Api-

Staph test (bioMerieux, Lyon, France). Bacterial

isolates were numbered 1 – 5 for S. epidermidis and

6 – 9 for S. aureus.

Bacterial DNA extraction

Bacteria were harvested by centrifuging 100 ml of

broth culture at 4000 g for 10 min. Cells were

resuspended in 45 ml H2O, 5 ml of lysostaphin

solution were added, and the samples were incu-

bated at 378C. After 10 min, 5 ml proteinase K

solution and 150 ml 0.1 M Tris-HCl (pH 7.5) were

added, and incubation proceeded for an additional

10 min. Samples were then heated for 5 min at

1008C (Arciola et al. 2001).

PCR amplification of icaA, icaD, and ica operon

(icaADBC)

Primers specific for icaA and icaD were used (Arciola

et al. 2001). The primers were synthesised by Iontek

(Istanbul, Turkey). For the detection of icaA, 50-TCTCTTGCAGGAGCAATCAA was used as the

forward primer (primer 1, corresponding to nucleo-

tides 1337 – 1356), and 50-TCAGGCACTAACAT-

CCAGCA was used as the reverse primer (primer 2,

corresponding to nucleotides 1505 – 1524). These two

primers include a 188-bp region. For detection of

icaD, 50-ATGGTCAAGCCCAGACAGAG was used

as the forward primer (primer 1, corresponding to

nucleotides 1963 – 1982), and 50-CGTGTTTTC-

AACATTTAATGCAA was used as the reverse

primer (primer 2, corresponding to nucleotides

2138 – 2160). These two primers include a 198-bp

region. PCR was performed in a DNA thermal cycler

in a 25-ml reaction mixture containing the above-

mentioned primers (1 mM each), together with 150 ng

extracted DNA, 100 mM dATP, dCTP, dGTP, and

dTTP, 1 U of Taq DNA polymerase, and buffer

(10 mM Tris-HCl [pH 9.0], 50 mM KCl, 0.1%

Triton X-100, and 2.5 mM MgCl2). A thermal step

program for icaA and icaD was used, which included

the following parameters: incubation at 948C for

5 min, followed by 50 cycles at 948C for 30 sec

(denaturation), 55.58C for 30 sec (annealing), 728Cfor 30 sec (extension), and 728C for 1 min after

conclusion of the 50 cycles. After the first 30 cycles, an

additional 1 U of Taq DNA polymerase was added.

After amplification, 10 ml PCR mixture were

analysed by agarose gel electrophoresis (2% agarose

in Tris-borate-EDTA). The MW marker kit pBR322

64 A. Karadenızlı et al.

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

DNA/BsuRI (HaeIII) Marker, 5 (Fermentas, MBI)

was used. The presence of the entire ica operon in

both S. epidermidis and S. aureus was checked by

amplifying a 2.7-kb gene product encompassing a

region of the icaADBC locus, as previously described

(McKenney et al. 1999). The PCR forward primer

TGCACTCAATGAGGGAATCA, corresponding

to nucleotides 409 – 428 of the icaA gene, was used;

the reverse primer AATCACTACCGGAAACA

GGC, complementary to nucleotides 3114 – 3133

of the icaC gene, was used. PCR was carried out

with platinum PCR Supermix and 200-nM primers.

DNA melting was performed at 958C for 30 s,

annealing was performed at 608C for 60 s, and

elongation was performed at 728C for 60 s. Amplified

DNA was visualised after separation in a 1% agarose

gel and staining with ethidium bromide.

Slime production

The tube adherence test was used to ensure that the

Staphylococcus strains had the ability to produce large

amounts of slime (Christensen et al. 1982). Briefly,

the strain was grown overnight in tryptic soy broth.

The tube was then emptied, stained with 0.1%

safranin for approximately 10 sec, and dried. The

tube was macroscopically examined for the presence

of a film on the inside walls. Strongly adherent

strains were chosen for further analysis. All slime-

producing strains were tested in duplicate. Produc-

tion of slime was also tested by cultivation of the

strains on Congo Red Agar (CRA) (Freeman et al.

1989).

Slime extraction by phenol

Cells were grown overnight at 378C in tryptic soy

broth. After being washed thrice with 0.9% NaCl,

they were resuspended in saline (*108 cells ml71).

Nutrient agar plates (11 cm diameter) containing 3%

casamino acids and 1% glucose were inoculated with

5 ml bacteria-saline suspension and incubated at

378C for 48 h. The liquid bacteria-slime mixture was

then collected in sterile conical tubes and vigorously

vortexed for 20 min to disrupt clots of slime. After

centrifugation at 12,000 g for 30 min, the super-

natant was dialysed against distilled water for 48 h at

48C, concentrated, and lyophilised. The crude slime

was subjected to phenol extraction and freeze-dried.

This material represented the phenol extracted slime

(PES) preparation (Sultan et al. 1997). Material was

also tested for carbohydrate content by alpha-

naphthol assay. A control sample was obtained in

the same manner as used for harvesting from the

slime-producing bacteria. The control extraction was

performed after incubating 5 ml sterile 0.9% NaCl

(without bacteria) on agar at 378C for 48 h.

FTIR spectroscopy and data analysis

The slime samples were ground and dried in a

MAXI dry lyo freeze drier for overnight. The

samples were then ground with potassium bromide,

a filling substance, at a ratio of 1 (sample):100 (KBr).

The powder was then compressed into a thin KBr

disk at 100 kg cm72 for 5 min. Spectral analysis was

performed using a BOMEM MB157 FTIR spectro-

meter (The Michelson Series, Bomem, Inc. Quebec,

Canada) equipped with a DTGS (deuterated trigly-

cine sulfate) detector. The sample compartment was

continuously purged with dry air to minimise

interference from water vapour and CO2. FTIR

spectra of samples were recorded in the 4000 –

400 cm71 region at room temperature. Four hun-

dred scans were taken for each interferogram at

4 cm71 resolutions. Each sample was scanned three

times, yielding identical results that were averaged

for further comparison. The GRAMS/32 soft-

ware programme (Galactic Industries Corporation,

Salem, NH, USA) was used to average these average

spectra, yielding the overall mean spectra for each

group. Following the FTIR spectroscopic measure-

ments, all spectral analyses were performed with

Win-Bomem Easy for Microsoft Windows Version

3.04 (Galactic Industries Corporation). The spec-

trum of the KBr pellet (pure KBr) was captured and

subtracted from all the samples interactively to yield

a slime sample spectrum. Baseline correction was

made and using the same software, Win-Bomem

Easy, the spectra were normalised to the 1588 peak,

which was prominent. Smoothed second derivatives

of the original spectra were then calculated using a

9-point Savitzky-Golay filter.

Hydrolysis of polysaccharides

Slime was taken into screw-cap tubes (136100 mm)

and 4 M HCl was added. After hydrolysis in a heating

block at 1008C for 2 h, the tubes were cooled, and the

acid was removed by evaporation with nitrogen gas

(Reddy et al. 1998). Hydrolysates were dried and

dissolved in 1 ml bidistilled water.

TLC

TLC was conducted using three ascents through a

solvent system of acetonitrile/water (90:10 v/v) on

Whatman K5 silica plates, running 5 ml of each

sample, an agar sample, and a control. After

hydrolysis, all samples and 1 mg agar powder were

dissolved in 1 ml double distilled water. Carbohy-

drates were visualised by dipping the plates into 5%

(v/v) H2SO4 in ethanol containing 0.5% (w/v) alpha-

naphthol, followed by heating at 1108C for 10 min

(Tanriseven & Dogan, 2002).

Classification of slime from staphylococci 65

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

Statistical and cluster analysis

Peak intensities of identical strains were compared

against the other strain S. aureus vs. S. epidermidis.

Statistical analyses were performed using the Mann-

Whitney U-tests. The construction of the dendo-

grams was performed executing Ward’s algorithm

implemented with the SPSS (version 11.5 for

Windows) option ‘‘Hierarchical cluster analysis’’

and selecting defined spectral regions of the second

derivative spectra as input data.

Results and discussion

Genetic analysis

The detection of genes governing the production of

EPS and, in particular, the icaA, icaC, and icaD

genes, is a rapid and accurate method for strain

characterisation. However, well-established meth-

ods, such as the CRA test, were necessary to confirm

phenotypic expression in the case of possible phase-

variant strains (Arciola et al. 2002). To eliminate

differences arising from the ica locus, attempts were

made to genetically characterise the ability of the two

strains to form slime. Both strains were positive for

the icaA and icaD genes and the entire ica operon

(2.7 kb); and both coloured CRA black (data not

shown). It was decided to use both assays to identify

positive slime formers since a recent report indicated

a lack of correlation between the presence of

icaA and icaD and slime production on CRA

(Vancraeynest et al. 2004).

Monosaccharide content analysis

Phenol-saline extracts from S. epidermidis and

S. aureus were hydrolysed and separated by TLC

using fucose, glucose, galactose, xylose, arabinose,

rhamnose, and mannose as reference markers. No

difference in profile between or within species was

detected with the naked eye, and monosaccharide

constituents yielded four visible bands with Rf values

0.54, 0.65, 0.85, and 0.95. The second of these

bands (Rf¼ 0.65) was identified as glucose, based on

monosaccharide markers. The Rf¼ 0.54 band was

consistent with fucose and xylose, but based on

comparison of the colour of the band with the

reference markers, xylose seemed more likely. No

band was visualised for the control (sterile 0.9%

NaCl from the surface of agar) (see Figure 1).

Earlier reports suggested that EPS is a complex

glycoconjugate containing protein and a variety of

monosaccharides including mannose, galactose, and

ribose (Peters et al. 1987). Ekstedt and Bernhard

(1973) isolated and partially purified slime from

S. aureus and found neutral sugars, among other

content, with galactose being the major component.

Using immunoelectron microscopy, Tojo et al. (1988)

characterised a specific polysaccharide capsular adhe-

sin that was a component of a capsular polysaccharide

in intact organisms, composed of a complex mixtures

of monosaccharides: galactose, glucosamine, and

galactosamine. Hussain et al. (1991) isolated protein-

free material containing glucose, glucosamine,

alanine, glycerol phosphate, and an additional uni-

dentified component from coagulase-negative Staphy-

lococcus. In studies of S. epidermidis slime, Ludwicka

et al. (1984) grew organisms under fluid on nutrient

agar (N-agar) supplemented with 3% (wt/vol) casami-

no acids and 1% (wt/vol) glucose and consistently

isolated a galactose-rich polysaccharide. Galactose and

galactosamine are known components of the phos-

phoglycoprotein (casein) used in the tryptic soy broth

(Jolles, 1972).

In an earlier investigation that included a sterile

control revealed that the galactose-rich product

previously referred to as slime was derived largely

from a contaminant of N-agar nutrient (Drewry et al.

1990). In the present study galactose was not

detected by TLC, although nutrient agar plates

containing 3% casamino acids were used, which is

the same agar condition used by Ludwicka et al.

(1984). Drewry et al. (1990) studied crude slime,

obtained by the growth of S. epidermidis (strain RP-

12) on submerged tryptic soy broth-agar, and found

that it contained 57% total carbohydrate and 1%

phosphorus. The major sugar component (galactose)

was accompanied by small amounts of glucose and

xylose and traces of mannose and ribose.

Figure 1. TLC of slime. Xylose and glucose were identified using monosaccharide markers (the sample band was identified as xylose based

on colour similarity to the marker xylose band). Bacterial isolates were numbered 1 – 5 for S. epidermidis and 6 – 9 for S. aureus. Fuc, fucose;

Glu, glucose; Gal, galactose; Xyl, xylose, Ara, arabinose; Rha, rhamnose; Mix, mixture; Con, control; Ag, agar.

66 A. Karadenızlı et al.

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

Further examination of the polymeric fractions

(test and control) showed the presence of glucose,

galactose, and galactosamine in all cases, together

with smaller amounts of arabinose and mannose.

These common components are likely derived from

the medium, possibly from glycans or glycoproteins

present in medium components. This is especially

true for products obtained from submerged solid

media (Drewry et al. 1990). In the present study,

Brain hearth infusion and N-agar were used, which

includes glucose, but the supernatant was dialysed

from the liquid bacteria-slime mixture after centri-

fugation against distilled water and slime was

obtained this way. Thus, free glucose was excluded

as a contaminant from the medium. Moreover, since

dialysing would not remove polymerised glucose

found in agar, a parallel sterile control was used.

TLC showed that the agar contained galactose (see

Figure 1), and the FTIR spectrum of the agar was

very different from that of the slime samples (see

Figure 2B). This confirmed the absence of glucose

contamination of the slime samples by polymerised

glucose in the agar.

Arvaniti et al. (1994) reported the isolation and

characterisation of a polysaccharide from the extra-

cellular slime layer of S. epidermidis. The polysac-

charide in the crude slime contained protein (11 –

20.5%), hexosamine (8 – 19%), the neutral sugars

(12.2 – 14%) galactose (0.13 – 1.2%), xylose (0.03 –

1.1%), glucose (1%), mannose (from trace – 0.8%),

and fucose (0.44 – 2.3%), phosphates (4 – 9.5%),

uronic acids (1 – 13%), and small amounts of sulfates

(0.5 – 3%) indicating the presence of carbohydrates

and protein/glycoproteins. They also revealed the

presence of one species of low-sulfated polysacchar-

ide with a relative molecular mass of 20 kDa.

Chemical analyses of the polysaccharide showed it

to be rich in glucosamine (46%) and neutral sugars

(30 – 34%). Glucose was the major neutral mono-

saccharide determined (60 – 65% of the total). It also

contained small amounts of fucose and xylose. In the

present study, a similar TLC profile was obtained for

all nine samples. One monosaccharide was identified

as glucose, and the other, which appeared to be the

most abundant of the four visible bands in all

analysed S. epidermidis and S. aureus strains, was

presumed to be xylose. This difference in findings

with respect to xylose may reflect the variability in

monosaccharide content within the same species.

In a study by Kotilainen et al. (1990), mannose,

galactose, glucose, and ribose were detected as the

main monosaccharide components of extracellular

extracts of S. epidermidis and Staphylococcus hominis

strains. Moreover, the mean relative concentrations

of these monosaccharides were essentially the same

for the different adherence phenotypes within the

species S. epidermidis. These results suggest the

absence of a causal connection between adherence

of coagulase-negative staphylococci and EPS pro-

duction of any of the four monosaccharides analysed

(Kotilainen et al. 1990).

FTIR characterisation of the slime

FTIR is a widely used technique in biofilm investiga-

tion (Nichols et al. 1985; Beech et al. 1999; Donlan

et al. 2004). A previous FTIR study showed that the

transition from surface adhesion to suspension

condition induced the disappearance of polysacchar-

ide fibers (Zeroual et al. 1994). This complex sugar

can also be located inside bacterial cells, in the form

of sugar reserves that are synthesised during incuba-

tion on solid medium (Geesey & White, 1990). In

the present study no structural difference between

the slime of S. aureus and S. epidermidis regarding the

parameters detected by FTIR spectra were found

and no shift was observed in the frequency values of

the bands (Toyran et al. 2004). The inclusion of

substantial amounts of NaCl (40.5 M) in the

medium stimulates the autolytic activity of S. aureus

(Cripps & Work, 1967; Madiraju et al. 1987) and

releases an extracellular DNA slime (Catlin &

Cunningam, 1958; Cripps & Work, 1967). DNA is

less likely to diffuse away from bacterial cells

attached to a solid surface than from those in a

liquid culture medium (Lorian, 1989). FTIR spec-

troscopy is also a sensitive technique for monitoring

nucleic acids (Jiang et al. 2004). In the current study,

the nucleic acid peak from the FTIR spectra of

phenol-extracted slime occurred at 1238 cm71 and

can be assigned to the asymmetric phosphate

(P¼¼O) stretching mode of PO2 in nucleic acids

(Nivens et al. 2001). However, no significant

variation was observed in this band. Although for

illustrative purposes spectra normalised with respect

to the specific selected band were used, all measure-

ments were performed using the corresponding

baseline-corrected original spectrum. The broad

complex region from 1200 – 900 cm71 has been

characterised in the literature as the region where

polysaccharides appear. Vibrations in this region are

characterised by C��OH stretching modes and

C��O��C, which is a glycosidic bond, C��O, ring-

stretching vibrations of carbohydrates (oligosacchar-

ides, polysaccharides, and alginate), C��O��P,

P��O��P in polysaccharides of cell wall, and P¼O

stretching (symmetrical) of PO2 in nucleic acids

(Bosch et al. 2005). Absorbances at 1100 –

1000 cm71 (C��OH and P��O stretching) and

1060 cm71 (C��OH stretching of alginate, which is

a polysaccharide extract) were previously observed in

biofilms produced by a mucoid Pseudomonas aerugi-

nosa isolate (Nivens et al. 2001). Figure 2 shows the

average spectra of S. aureus strains (dashed line) and

Classification of slime from staphylococci 67

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

S. epidermidis strains (connected line) in the whole

spectrum (A), in the 1200 – 900 cm71 polysacchar-

ide region (B), and in the 1060 – 1000 cm71 region

of the second derivative spectra (C). Recorded

spectra showed a characteristic triple peak centered

at 1030 cm71, which is typical for polysaccharides

that clearly differ within this region, reflecting

molecular differences (Hoiczyk, 1998). The control

(without bacteria, 5 ml sterile of 0.9% NaCl

obtained by using sterile agar) and the biofilm-

negative mucoid P. aeruginosa, S. aureus, and

S. epidermidis isolates produced no lyophilised

material to visualise FTIR spectra.

Classification of slime produced by S. aureus and

S. epidermidis

Based on the second derivative FTIR spectra from

the normalised ones and the narrow spectral ranges

of 1060 – 1030 cm71, 1060 – 1020 cm71, 1060 –

1000 cm71, 1050 – 1000 cm71, 1030 – 1000 cm71,

and 1050 – 970 cm71, a distinct clustering scheme

was obtained that discriminated between the slimes

produced by S. epidermidis and S. aureus. FTIR

spectra were used as an input for hierarchical

clustering. The spectral information contained in

these spectral ranges was used to calculate the

distance matrix as an input for cluster analysis,

which resulted in the dendogram shown in Figure 3.

Although the spectral range 1000 – 1060 cm71 was

given as an example, cluster analysis performed

with the second derivative spectra considering the

spectral ranges 970 – 1050 cm71, 1000 – 1030 cm71,

1000 – 1050 cm71, 1020 – 1060 cm71, and 1030 –

1060 cm71 with the same grouping also revealed

two clusters (data not shown). Nine phenol-

extracted slimes were classified into two clusters.

For S. aureus, three of four (75%) were grouped

correctly, and for S. epidermidis, four of five (80%)

were grouped correctly. TLC was unable to differ-

entiate the samples of S. aureus and S. epidermidis. In

slime structure, a 1 – 6 glycosidic and b 1 – 3 bonds

are known to exist in linkage and branching,

respectively (Koolman & Rohm, 1996). The FTIR

approach has considerable potential to provide iden-

tifications for species-level classification of various

Figure 2. (A) Average slime spectra of S. aureus (- - - -) and S. epidermidis (_____) strains in the whole spectrum, 400 – 4000 cm71 (spectra

were normalised to the 1588 peak). Arrow¼ the nucleic acid peak at 1238 cm71. (B) Polysaccharide region, 1200 – 900 cm71; the vertical

lines show the characteristic triple peak centered at 1030 cm71. Agar medium is also shown. (C) The second derivative spectra in 1000 –

1060 cm71 region where cluster analysis was applied; AU, arbitrary units.

68 A. Karadenızlı et al.

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

bacteria, including S. aureus and S. epidermidis, and

hence for epidemiological and screening purposes

(Ngo-Thi et al. 2003).

Cluster analysis is a tool for categorising similar

objects and discovers structures in data without

explaining why these structures exist. Ward’s Algo-

rithm is a commonly used procedure for forming

hierarchical groups of mutually exclusive subsets and

can be applied to cluster any data set for which a

distance or similarity measure is available. The

combined distance of horizontal lines indicates

relative similarity between slimes. The shorter the

distance, the greater the number of shared simila-

rities; the longer the distance, the fewer the number

of shared similarities.

The proposed novel method differentiates between

the different samples, despite their FTIR spectra

being quite similar. A narrow region in the poly-

saccharide spectral range was used as the input for

cluster analysis, yielding two different groups. Other

regions of the spectra investigated could not differ-

entiate the two groups. Since S. aureus and S.

epidermidis were grown in the same medium, the

differences in their FTIR spectra cannot be a result

of medium contamination. Thus, the FTIR techni-

que can be useful for classifications at not only the

species level but also the slime level. Identifying the

bacterial strain from slime scraped off the biomaterial

surface without culturing will be advantageous,

especially for nosocomial infections caused by

pathogenic strains.

Conclusion

EPS may comprise up to 98% of biofilm volume

(Fleming et al. 2000). In the present study, the

structure of slime from S. aureus was differentiated

from that from S. epidermidis in a manner not

classified by other methods and examination of slime

with FTIR enabled the differentiation of closely

related strains.

The polysaccharide regions from S. aureus and

S. epidermidis were reported to be distinct. The

findings demonstrate that polysaccharide content

plays a major role in slime structure classification.

The approach taken represents a powerful diagnostic

method for determining bacterial strain based on

slime analysis and a promising method for screening

isolates for their ability to produce drug-resistant

biofilms.

At least three mechanisms have been proposed to

account for the increased resistance of biofilms to

antimicrobial agents. First, the biofilm glycocalyx

prevents the perfusion of antimicrobial agents.

Secondly, the nearly dormant growth pattern of

bacterial populations within the biofilm render them

indifferent to antibiotic activity. Thirdly, increased

resistance occurs due to adverse effects of the biofilm

microenvironment on antimicrobial activity (Dunne,

2002). A better understanding of the potential

mechanisms for biofilm resistance is of primary

importance. As one of the parameters that cause

stickiness on biomaterial surfaces, polysaccharide

content may gain new relevance in Staphylococcus

pathogenesis. Understanding the factors responsible

for differences in slime content and structure may

have importance in guiding therapy and is a target for

further investigation.

Acknowledgements

This work was supported by the Research Fund of

the Kocaeli University (Grant 2002/9 and 2002/52)

and former ‘‘Troika’’. The authors thank Dr Feride

Severcan for providing the FTIR Spectroscopy

facilities and Dr Ali Sazci for critically reading the

manuscript.

References

Allen MS, Welch KT, Prebyl BS, Baker DC, Meyers AJ,

Sayler GS. 2004. Analysis and glycosyl composition of

the exopolysaccharide isolated from the floc-forming waste-

water bacterium Thauera sp. MZ1T. Environ Microbiol 6:

780 – 790.

Ammendolia MG, Di Rosa R, Montanaro L, Arciola CR,

Baldassarri L. 1999. Slime production and expression of

the slime-associated antigen by staphylococcal clinical isolates.

J Clin Microbiol 37:3235 – 3238.

An YH, Friedmann RJ. 1998. Concise review of mechanisms of

bacterial adhesion to biomaterial surfaces. J Biomed Mater Res

43:338 – 348.

Arciola CR, Baldassarri L, Montanaro L. 2001. Presence of icaA

and icaD genes and slime production in a collection of

staphylococcal strains from catheter-associated infections.

J Clin Microbiol 39:2151 – 2156.

Arciola CR, Campoccia D, Montanaro L. 2002. Detection of

biofilm-forming strains of Staphylococcus epidermidis and S

aureus. Expert Rev Mol Diagn 2:478 – 484.

Figure 3. Dendogram showing the classification of slime spectra

for S. aureus and S. epidermidis using hierarchical cluster analysis.

Cluster analysis was performed with the second derivative spectra

considering the spectral range 1000 – 1060 cm71. Ward’s

algorithm was applied to construct the dendogram.

Classification of slime from staphylococci 69

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

Arciola CR, Montanaro L, Baldassarri L, Borsetti E,

Cavedagna D, Donati ME. 1999. Slime production by

staphylococci isolated from prosthesis-associated infection.

New Microbiol 22:337 – 341.

Arvaniti A, Karamanos NK, Dimitracopoulos G, Anastassiou ED.

1994. Isolation and characterization of a novel 20-kDa sulfated

polysaccharide from the extracellular slime layer of Staphylo-

coccus epidermidis. Arch Biochem Biophys 308:432 – 438.

Barth E, Myrvik QM, Wagner W, Gristina AG. 1989. In vitro and

in vivo comparative colonization of Staphylococcus aureus and

Staphylococcus epidermidis on orthopaedic implant materials.

Biomaterials 10:325 – 328.

Beech I, Hanjagsit L, Kalaji M, Neal AL, Zinkevich V. 1999.

Chemical and structural characterization of exopolymers

produced by Pseudomonas sp. NCIMB 2021 in continuous

culture. Microbiology 145:1491 – 1497.

Bosch A, Serra D, Prieto C, Schmitt J, Naumann D, Yantorno O.

2005. Characterization of Bordatella pertussis growing as biofilm

by chemical analysis and FT-IR spectroscopy. Appl Micobiol

Biotechnol 16:1 – 12.

Catlin BW, Cunningam LS. 1958. Studies of extracellular and

intracellular bacterial deoxyribonucleic acids. J Gen Microbiol

19:522 – 539.

Christensen GD, Baldassarri L, Simpson WA. 1994. In:

Bisno AL, Waldvogel FA, editors. Infections associated with

indwelling medical devices. Washington, DC: American

Society for Microbiology. 2nd ed. pp 45 – 78.

Christensen GD, Simpson WA, Bisno AL, Beachey EH. 1982.

Adherence of slime-producing Staphyloccus epidermidis to

smooth surfaces. Infect Immun 37:318 – 326.

Costerton JW, Cheng KJ, Geesey GG, Ladd TI, Nickel JC,

Dasgupta M, Marrie TJ. 1987. Bacterial biofilms in nature and

disease. Annu Rev Microbiol 41:435 – 464.

Crampton SE, Gerke C, Schnell NF, Nichols WW, Gotz F. 1999.

The intercellular adhesion (ica) locus is present in Staphylo-

coccus aureus and is required for biofilm formation. Infect

Immun 67:5427 – 5433.

Cripps RE, Work E. 1967. The accumulation of extracellular

macromolecules by Staphylococcus aureus grown in the presence

of sodium chloride and glucose. J Gen Microbiol 49:127 – 137.

Davies DG, Chakrabarty AM, Geesey GG. 1993. Exoploysacchar-

ide production in biofilms: substratum activation of alginate

gene expression by Pseudomonas aeruginosa. Environ Microbiol

59:1181 – 1186.

Donlan RM, Piede JA, Heyes CD, Sanii L, Murga R, Edmonds P,

El-Sayed I, El-Sayed MA. 2004. Model system for growing and

quantifying Streptococcus pneumoniae biofilms in situ and in real

time. Appl Environ Microbiol 70:4980 – 4988.

DrewryDT,GalbraithL,WilkinsonBJ,WilkinsonSG.1990.Staphy-

lococcal slime: a cautionary tale. J Clin Microbiol 28:1292 – 1296.

Dunne WM Jr. 2002. Bacterial adhesion: seen any good biofilms

lately? Clin Microbiol Rev 15:155 – 166.

Ekstedt RD, Bernhard JM. 1973. Preparation and characterization

of a slime layer material produced by Staphylococcus aureus. Proc

Soc Exp Biol Med 142:86 – 91.

Feigin RD, Kline MW, Hyatt SR, Ford III KL. 1992. Otitis

media. In: Feigin RD, Cherry JD, editors. Textbook of

pediatric infectious diseases. 3rd ed, Vol. 1. Philadelphia:

W. B. Saunders Co. pp 174 – 189.

Fleming HC, Wingender J, Griebe T, Mayer C. 2000. Physico-

chemical properties of biofilms. In: Evans LV, editor. Biofilms:

recent advances in their study and control. Amsterdam, The

Netherlands: Harwood Academic Publishers. pp 19 – 34.

Foster TJ, McDevitt D. 1994. Molecular basis of adherence of

staphylococci to biomaterials. In: Bisno AL, Waldvogel FA,

editors. Infection associated with indwelling medical devices.

Washington, DC: American Society for Microbiology. 2nd ed.

pp 31 – 43.

Freeman DJ, Falkiner FR, Keane CT. 1989. New method for

detecting slime production by coagulase-negative staphylococci.

J Clin Pathol 42:872 – 874.

Frere JM, Dubus A, Fonze E. 1999. Pathogens old, new and

revived. Nat Biotechnol 17:717 – 718.

Geesey GG, White DC. 1990. Determination of bacterial growth

and activity at solid-liquid interfaces. Annu Rev Microbiol

44:579 – 602.

Gerke C, Kraft A, Sussmuth R, Schweitzer O, Gotz F. 1998.

Characterization of the N-acetylglucosaminyltransferase activity

involved in the biosynthesis of the Staphylococcus epidermidis

polysaccharide intercellular adhesin. J Biol Chem 273:18586 –

18593.

Gotz F. 2002. Staphylococcus and biofilms. Mol Microbiol

43:1367 – 1378.

Hoiczyk E. 1998. Structural and biochemical analysis of the sheath

of Phormidium uncinatum. J Bacteriol 180:3923 – 3932.

Hoyle BD, Williams LJ, Costerton JW. 1993. Production of

mucoid exopolysaccharide during development of Pseudomonas

aeruginosa biofilms. Infect Immun 61:777 – 780.

Huebner J, Goldmann DA. 1999. Coagulase-negative staphylo-

cocci: role as pathogens. Annu Rev Med 50:223 – 236.

Hussain M, Hastings JG, White PJ. 1991. Isolation and composition

of the extracellular slime made by coagulase-negative staphylo-

cocci in a chemically defined medium. J Infect Dis 163:534 – 541.

Jolles P. 1972. Casein. In: Gottschalk A, editor. Glycoproteins:

their composition, structure and function. Part B. Amsterdam:

Elsevier Biomedical Press. pp 782 – 809.

Jiang W, Saxena A, Song B, Ward BB, Beveridge TJ, Myneni

SCB. 2004. Elucidation of functional groups on Gram-positive

and Gram-negative bacterial surfaces using infrared spectro-

scopy. Langmuir 20:11433 – 11442.

Koolman J, Rohm K-H. 1996. Disaccharides and polysaccharides.

In: Koolman J, Rohm K-H, editors. Color atlas of biochemis-

try. Stuttgart: Georg Thieme Verlag. p 37.

Kotilainen P, Oksman P, Viljanen MK, Nikoskelainen J, Huovinen

P. 1990. Analysis of the relationship between bacterial

adherence and extracellular production of mannose, galactose,

glucose and ribose in Staphylococcus epidermidis and Staphylo-

coccus hominis. Eur J Clin Microbiol Infect Dis 9:873 – 879.

Lorian V. 1989. In vitro simulation of in vivo conditions: physical

state of the culture medium. J Clin Microbiol 27:2403 – 2406.

Ludwicka A, Uhlenbruck G, Peters G, Seng PN, Gray ED,

Jeljaszewicz J, Pulverer G. 1984. Investigation on extracellular

slime substance produced by Staphylococcus epidermidis. Zen-

tralbl Bakteriol Mikrobiol Hyg [A] 258:256 – 267.

MacKintosh EE, Patel JD, Marchant RE, Anderson JM. 2006.

Effects of biomaterial surface chemistry on the adhesion and

biofilm formation of Staphylococcus epidermidis in vitro. J Biomed

Mater Res A 78:836 – 842.

Madiraju MVVS, Brunner DP, Wilkinson BJ. 1987. Effects of

temperature, NaCl, and methicillin on penicillin-binding

proteins, growth, peptidoglycan synthesis, and autolysis in

methicillin-resistant Staphylococcus aureus. Antimicrob Agents

Chemother 31:1727 – 1733.

McKenney D, Pouliot KL, Wang Y, Murthy V, Ulrich M, Doring

G, Lee JC, Goldmann DA, Pier GB. 1999. Broadly protective

vaccine for Staphylococcus aureus based on an in vivo-expressed

antigen. Science 284:1523 – 1527.

Montanaro L, Arciola CR. 2000. Studying bacterial adhesion

to irregular or porous surfaces. In: An YH, Friedmann RJ,

Totowa NJ, editors. Handbook of bacterial adhesion: princi-

ples, methods and applications. Totowa, NJ: Humana Press

Inc. pp 331 – 343.

Montanaro L, Arciola CR, Borsetti E, Brigotti M, Baldassarri L.

1998. A polymerase chain reaction (PCR) method for the

identification of collagen adhesin gene (cna) in Staphylococcus-

induced prosthesis infections. New Microbiol 21:359 – 363.

70 A. Karadenızlı et al.

Dow

nloa

ded

By: [

ANKO

S 20

07 O

RD

ER C

onso

rtium

] At:

07:4

8 3

May

200

7

Ngo-Thi NA, Kirschner C, Naumann D. 2003. Characterization

and identification of microorganisms by FT-IR microspectro-

metry. J Mol Struct 661:371 – 380.

Nichols PD, Henson JM, Guckert JB, Nivens DE, White DC.

1985. Fourier transform-infrared spectroscopic methods for

microbial ecology: analysis of bacteria, bacteria-polymer mix-

tures and biofilms. J Microbiol Methods 4:79 – 94.

Nivens DE, Ohman DE, Williams J, Franklin MJ. 2001. Role of

alginate and its O acetylation in formation of Pseudomonas

aeruginosa microcolonies and biofilms. J Bacteriol 183:1047 –

1057.

Peters G, Schumacher-Perdreau F, Jansen B, Bey M, Pulverer G.

1987. In: Pulverer G, Quie PG, editors. Pathogenicity and

clinical significance of coagulase-negative staphylococci.

Stuttgart: Fischer-Verlag. pp 15 – 31.

Reddy GP, Hayat U, Xu Q, Reddy KV, Wang Y, Chiu KW,

Morris JG Jr, Bush CA. 1998. Structure determination of the

capsular polysaccharide from Vibrio vulnificus strain 6353. Eur J

Biochem 255:279 – 288.

Rupp ME, Archer GD. 1994. Coagulase-negative staphylococci:

pathogens associated with medical progress. Clin Infect Dis

19:231 – 245.

Severcan F, Sahin I, Kazanci N. 2005. Melatonin strongly

interacts with zwitterionic model membranes – evidence from

Fourier transform infrared spectroscopy and differential scan-

ning calorimetry. Biochim Biophys Acta 1668:215 – 222.

Severcan F, Toyran N, Kaptan N, Turan B. 2000. Fourier

transform infrared study of the effect of diabetes on rat liver and

heart tissues in the C-H region. Talanta 53:55 – 59.

Sultan N, Caglar K, Aybay C, Imir T. 1997. Effect of a

Staphylococcus epidermidis-extracted slime factor on human

natural killer cell activity. Microbios 91:89 – 95.

Tanriseven A, Dogan S. 2002. A novel method for the

immobilization of b-galactosidase. Proc Biochem 38:27 – 30.

Tojo M, Yamashita N, Goldmann DA, Pier GB. 1988. Isolation

and characterization of a capsular polysaccharide adhesin from

Staphylococcus epidermidis. J Infect Dis 157:713 – 722.

Toyran N, Zorlu F, Donmez G, Oge K, Severcan F. 2004.

Chronic hypoperfusion alters the content and structure of

proteins and lipids of rat brain homogenates: a Fourier trans-

form infrared spectroscopy study. Eur Biophys J 33:549 –

554.

Vancraeynest D, Hermans K, Haesebrouck F. 2004. Genotypic

and phenotypic screening of high and low virulence Staphylo-

coccus aureus isolates from rabbits for biofilm formation and

MSCRAMMs. Vet Microbiol 103:241 – 247.

Vann WF, Moreau M, Sutton A, Byrd RA, Karakawa WW. 1988.

Structure and immunochemistry of Staphylococcus aureus

capsular polysaccharides. In: Horowitz MA, editor. Bacteria-

host-cell interaction. UCLA Symposia on Molecular and

Cellular Biology Series, Vol. 64. New York: Alan R. Liss,

Inc. pp 187 – 198.

Vaudaux P, Yasuda H, Velazco MI, Huggler E, Ratti I, Waldvogel

FA, Lew DP, Proctor A. 1990. Role of host and bacterial

factors in modulating staphylococcal adhesion to implanted

polymer surfaces. J Biomater Appl 5:134 – 153.

Wilkinson BJ. 1983. Staphylococcal capsules and slime. In:

Easmon CSF, Adlam C, editors. Staphylococci and staphylo-

coccal infections, Vol. 2. New York: Academic Press, Inc.

pp 481 – 523.

Zeroual W, Choisy C, Doglia SM, Bobichon H, Angiboust JF,

Manfait M. 1994. Monitoring of bacterial growth and structural

analysis as probed by FT-IR spectroscopy. Biochim Biophys

Acta 1222:171 – 178.

Classification of slime from staphylococci 71