Adsorbed poly(ethyleneoxide)–poly(propyleneoxide)
copolymers on synthetic surfaces: Spectroscopy and
microscopy of polymer structures and effects on adhesion
of skin-borne bacteria
Lorraine H. Marsh,1 Mark Coke,2 Peter W. Dettmar,2 Richard J. Ewen,3 Michael Havler,2 Thomas G. Nevell,1
John D. Smart,1 James R. Smith,1 Barry Timmins,2 John Tsibouklis,1 Cameron Alexander1
1
School of Pharmacy and Biomedical Science, University of Portsmouth, White Swan Road,
Portsmouth PO1 2DT, United Kingdom
2
Reckitt Benckiser Healthcare (UK) Limited, Dansom Lane, Hull, HU8 7DS, United Kingdom
3
Faculty of Applied Sciences, University of the West of England, Frenchay Campus, Coldharbour Lane,
Bristol BS16 1QY, United Kingdom
Received 15 February 2001; revised 12 October 2001; accepted 28 November 2001
Abstract: Poly(ethyleneoxide)–copoly(propyleneoxide)
(PEO-PPO) polymer coatings were evaluated for their resistance to the attachment of the marker organism Serratia marcescens and the skin-borne bacteria Staphylococcus epidermidis.
The copolymers were adsorbed onto poly(styrene) films—
chosen as simplified physicochemical models of skin surfaces—and their surface characteristics probed by contact
angle goniometry, attenuated total reflectance–Fourier
transform infrared (ATR-FTIR), atomic force microscopy
(AFM), and X-ray photoelectron spectroscopy (XPS). These
functional surfaces were then presented to microbial cultures, bacterial attachment was assessed by fluorescence microscopy and AFM, and the structures of the polymer films
examined again spectroscopically. Surface characterization
data suggest that the adsorbed copolymer was partially re-
tained at the surface and resisted bacterial attachment for 24
h. Quantitative evaluation of cell attachment was carried out
by scintillation counting of 14C-labeled microorganisms in
conjunction with plate counts. The results show that a
densely packed layer of PEO-PPO copolymer can reduce
attachment of skin commensals by an order of magnitude,
even when the coating is applied by a simple adsorptive
process. The work supports the hypothesis that adhesion of
microorganisms to biological substrates can be reduced if a
pretreatment with an appropriate copolymer can be effected
in vivo. © 2002 Wiley Periodicals, Inc. J Biomed Mater Res 61:
641–652, 2002
INTRODUCTION
venting the attachment of biopolymers and cells has
been investigated in great detail.2 The pioneering
studies of Harris,3–5 Andrade,6,7 and Merrill8,9 using
poly(ethyleneglycol) (PEG) derivatives, and more recent work by the groups of Marchant,10,11 Park,12 Davies,13,14 and Whitesides,15,16 have shown that biomimetic hydrophilic surfaces in particular are highly effective in preventing protein and cell adsorption.
However, the quest for experimentally straightforward and easily practicable methods by which the attachment of skin commensal microorganisms to preformed or biological surfaces may be prevented has
attracted less attention.17 For example, a facile nonbiocide treatment process that can protect skin surfaces
from pathogen attachment and thus prevent wound
invasion or medical implant colonization is of both
clinical and economic importance, but has yet to be
fully realized. This can be attributed partly to the fact
The preparation of biocompatible synthetic surfaces
and fouling-resistant coatings is of great interest, and
such materials could find many applications in medicine, industry, and the home.1 As a consequence, large
numbers of potentially fouling-resistant materials
have been developed and their performance in preThe author, or one or more of the authors, has received or
will receive remuneration or other prerequisites for personal
or professional use from a commercial or industrial agent in
direct or indirect relationship to their authorship.
Correspondence to: C. Alexander; e-mail: cameron.
[email protected]
Contract grant sponsor: Reckitt Benckiser Healthcare and
BBSRC
© 2002 Wiley Periodicals, Inc.
Key words: biocompatible polymers; bacterial adsorption;
biofouling; surface analysis; Poly(ethyleneoxide)–
copoly(propyleneoxide) copolymers
642
that the relationships between the chemical functionality of a surface and the mechanisms and/or extent of
bioadhesion are inherently highly complex18–20 and
also because methods derived for generating antifouling surfaces on existing substrates often require aggressive reagents or conditions.21,22 As a result, suitable coating regimes are not always accessible for the
treatment of natural materials or delicate substrates
prior to their introduction into biological environments. Nonetheless, irrespective of whether a substrate is synthetic or biological, it is likely that the
molecular and submolecular level structures adopted
by an applied coating at the surface are vital in regulating the attachment of biopolymers and microorganisms. 23,24 These surface structures are of course
strongly influenced by the experimental methods used
to derive them, and it is this surface preparation that is
of key importance where practical applications are envisaged or in the treatment of biological substrates.
Whilst the “steric stabilization” effects of poly(ethyleneoxide) coatings in general have been very widely
studied,25 it is less clear as to whether these and other
commercially available hydrophilic block copolymers
can form suitably structured surfaces without elaborate coating methods or retain their antifouling properties for prolonged periods (up to 24 h) during challenge with bacterial cultures. The successful demonstration of easily applicable and durable antifouling
coatings, especially against microorganisms that can
be present on human skin such as Serratia marcescens,
and Staphylococcus epidermidis and that are implicated
in common infection,26 would be of obvious benefit in
household applications and in medicine.
The aim of the work presented here was therefore to
devise a method for preparing hydrophilic antifouling
coatings (a) that was experimentally simple to carry
out, (b) that could be performed on easily characterizable surfaces possessing properties similar to those
of human skin cells, and (c) that would generate structured polymer surfaces that resisted the attachment of
bacteria.
We report here that a facile dip-coating process, using appropriate commercially available synthetic
polymers in aqueous solutions, can give rise to the
surface-displayed features and macromolecular orientations necessary to bestow resistance to bacterial attachment. We have used contact angle goniometry,
fluorescence spectroscopy, Fourier transform infrared
(FTIR), atomic force microscopy (AFM), and X-ray
photoelectron spectroscopy (XPS) experiments to
probe the structures generated on model surfaces by
dip-coating poly(ethyleneoxide)–copoly(propyleneoxide) block copolymers (PEO-PPO) and have
used these techniques in conjunction with fluorescence microscopy and bacterial radiolabeling experiments to confirm that densely packed and pseudocrystalline domains are required for good antifouling
MARSH ET AL.
performance. Furthermore, the study shows that polymer domains produced using the simple dip-coating
process remain at least partially on model skin-mimic
surfaces even after incubation with clinically relevant
bacterial cultures, and suggests that a similar methodology may be used to protect human skin surfaces
from bacterial colonization if the copolymer adsorption can be controlled in vivo.
EXPERIMENTAL
Materials and methods
Chemicals
Standard reagents and chemicals were purchased from
Fisher Scientific or Aldrich, and used as received. Poly(ethylene oxide)–copoly(propylene oxide) (PEO-PPO) copolymers (Pluronics™) were obtained from BASF (see Fig. 1).
Poly(methylmethacrylate) sheets (PMMA) (Goodrich) were
cut into samples of nominal dimensions 10 × 20 mm, carefully wiped with butanone [high performance liquid chromatography (HPLC) grade], thoroughly rinsed with copious
quantities of double distilled water, and then dried in an
oven at 60°C. Poly(dimethylsiloxane) (PDMS) was prepared
by reaction of Dow Corning 734 elastomer with ethyltriacetoxysilane and methyltriacetoxysilane. 27 The resultant
crosslinked PDMS gel samples were cut to size and mounted
on a microscope slide. D-(U-14C)-glucose was obtained from
Amersham Life Sciences (Amersham, UK). Dye-labeled
PEO-PPO was prepared by reaction of fluorescein isothiocyanate (FITC) (10 mg) with Pluronic F127 (1.0 g) in anhydrous dimethylformamide (DMF) (10 mL) and triethylamine
(1 mL) for 24 h, followed by exhaustive dialysis (4kDa cutoff,
15 × 1000 mL) in double-distilled water. When no further
FITC could be detected in the dialysate, the labeled polymer
was recovered by lyophilization. Purification of solvents for
preparative chemistry was by standard methods.28 For contact angle goniometry, double-distilled water (surface tension 72.8 mN m−1 at 20.0°C), diiodomethane (Aldrich >99%,
surface tension 50.7 mN m−1 at 20.0°C) and ethylene glycol
(Aldrich 99.8%, anhydrous, surface tension 48.0 mN m−1 at
20.0°C) were used.
Preparation of glass surfaces and polymer coatings
Silica glass slides (Chance Propper, UK) were cleaned
with 5M sodium hydroxide solution, rinsed in deionized
Figure 1. Structures of Pluronic™ copolymers. F127: x = 99,
y = 67, z = 99 (31% PPO, 69% PEO). Molecular weight ∼
12,600.
ADSORBED PEO-PPO COPOLYMERS ON SYNTHETIC SURFACES
water (10 × 100 mL), dried, immersed in fresh ammonium
persulfate/sulfuric acid (1% w/v) or chromic acid, washed
again (10 × 100 mL) in deionized water and ethanol, then
dried at 80°C for 24 h. The cleaned glass slides were silanized by immersion in a solution containing 0.5% w/v dimethyldichlorosilane in octamethylcyclotetramethylsiloxane, before being washed in hexane (3 × 50 mL) and
acetone (3 × 50 mL), and then dried. Poly(styrene) (PS) films
were applied by dipping (10 min) the hydrophobized slides
in a solution of poly(styrene) (5% w/v) in toluene. PEO-PPO
polymer coatings were subsequently deposited on the surfaces by dip coating PS–glass slides in aqueous solutions of
Pluronic copolymer (1–2% w/v in double-distilled water)
followed by drying under a flow of dry nitrogen.
Growth and labeling of microorganisms
Bacterial cultures were maintained on nutrient agar (CM3
Oxoid) at 4°C. Stock cultures of Serratia marcescens and
Staphylococcus epidermidis were grown statically (overnight,
37°C) in nutrient broth (Oxoid), and aliquots (100 ␮L) of
each culture in sterile Eppendorf tubes were stored at −70°C
prior to use. For fluorescence microscopy the bacterial cultures were prestained with ethidium bromide solution (10
mg mL−1, 1 mL) incubated for 3 h, then centrifuged (5 min,
3000 rpm). The supernatants were removed and pellets of
microorganisms were resuspended in broth (10 mL) by vortex mixing (30 s). To obtain 14C-glucose-labeled bacteria, nutrient broth (6 mL) was first inoculated with thawed stock
culture (50 ␮L), and an aliquot (100 ␮L) containing 20 ␮Ci of
D-(U-14C)-glucose from a 200 ␮Ci mL−1 stock solution was
added. The culture was then incubated (37°C, 24 h) statically. To determine absolute cell numbers, aliquots (200 ␮L)
of unlabeled organisms were enumerated via a serial dilution method. The absolute cell numbers determined in this
way were compared with scintillation counts from equivalent aliquots (200 ␮L) of radiolabeled bacteria.
643
sis system was used. Drops of liquid of known volume (1–4
␮L) were applied from a microsyringe to the surface of the
test material through a small port at the top of the cell: to
avoid cross-contamination of liquids, a dedicated microsyringe was used for each diagnostic liquid. The precision of
measurement was ±0.5°.
Atomic force microscopy
AFM studies were performed in air under ambient conditions, using a Discoverer TopoMetrix TMX2000 scanning
probe microscope (SPM), which was mounted on a custombuilt mass/spring antivibration rig with a lateral natural
frequency of 0.40 Hz and a vertical natural frequency of 0.52
Hz. “V”-shaped silicon nitride cantilevers of length 200 ␮m
and nominal spring constant (k) 0.032 N.m−1, bearing an
integrated standard profile tip (part no. 1520-00 ThermoMicroscopes, Santa Clara, CA) were used.
Infrared spectroscopy
Infrared spectra were obtained employing a Perkin Elmer
Paragon 1000 FT-IR spectrophotometer, in both transmittance and ATR modes, at a resolution of 4 cm−1.
UV spectroscopy
A Cecil CE1010 spectrophotometer (Cambridge, UK) was
used.
X-ray photoelectron spectroscopy
In vitro adsorption studies
Bacterial cultures (107–108 CFU ⭈ mL−1), either radiolabeled or stained with ethidium bromide (6 mL) were transferred to centrifuge tubes and centrifuged (8000 rpm, 10
min), washed twice in sterile 3-[N-morpholino]propanesulfonic acid (MOPS) (50 mM, pH 7.0, 6 mL), and resuspended
in MOPS (6 mL). Aliquots (200 ␮L) of the suspended bacteria
were transferred to sterile MOPS (10 mL) in capped bottles.
Glass test surfaces (1 cm2), with and without polymer coatings, were then placed in the capped bottles and incubated,
with gentle shaking (40 rpm), for 24 h at 37°C. The slides
were rinsed in sterile MOPS (10 mL), and then examined by
fluorescence microscopy or transferred to scintillation vials
containing Ultima Gold scintillant cocktail (5 mL). All experiments were conducted in triplicate.
Instrumentation
XPS data were obtained using a VG Scientific ESCALAB
Mk. II employing a nonmonochromatized Al K source
(1486.6 eV) operating at a power of 125 W and a take-off
angle of 75°. The analyzer was operated at constant pass
energy of 20 eV. Line shape analysis was performed on each
peak, and atomic percentages were calculated from the peak
areas using standard atomic sensitivity factors.29
Fluorescence microscopy
A Vickers Epifluorescent microscope, with a broad band
mercury lamp fitted with a green excitation filter system,
was employed.
Scintillation counting
Contact angle goniometry
A Kruss G10 contact angle measuring system equipped
with a sealed sample chamber, and automated image analy-
A Packard Tri-Carb 1900TR Liquid Scintillation Analyser
was used (Packard Instrument Company, Meriden, CT,
USA).
644
MARSH ET AL.
RESULTS AND DISCUSSION
The primary target was to attach hydrophilic copolymers to model surfaces without complex chemical
grafting procedures yet still generate an entropic barrier to adhesion of skin-borne bacteria. However, it
was also important that we were able to coat the polymers on surfaces that were close mimics of human
skin in terms of overall chemical characteristics, but
that were amenable to repeated physical, spectroscopic, and microscopic analysis. In view of the difficulties (cell availability, infective risk, experimental
variation, and ethical constraints) of working with human skin samples, it was first necessary to develop a
convenient in vitro model that possessed similar physicochemical properties to human skin, but that was
readily accessible to conventional surface characterization techniques such as IR spectroscopy and AFM.
Thus we required a model-skin surface that was flat,
spectroscopically transparent relative to adsorbed hydrophilic copolymers and yet of predominantly the same
adsorptive properties as human epidermal cells. Synthetic polymer models for skin have been derived previously using contact angles as a probe of adsorptive
behavior,30 and surface energy components of skin
have also been used to evaluate barrier properties and
protective function.31 However, there are also limitations to the use of skin models in general,32,33 and in
using human skin in vivo owing to individual subject
variations and ethical constraints. As bacterial adsorption is inherently highly complex, it was necessary to
work with the simplest model systems possible before
embarking on detailed in vivo studies. In addition, the
biochemistry and physical hierarchy of skin are both
exceedingly complex and intrinsically dynamic; nevertheless, the initial adsorption behavior of synthetic
macromolecules onto most surfaces is largely a function of properties such as hydrophilicity/hydrophobicity and overall surface energy.34 This is because
for hydrophobic substrates, such as the outermost layers of skin (the strateum corneum and keratinocytes
are water resistant and essentially hydrophobic under
normal conditions35), thermodynamic reasons dictate
that adsorption of amphiphilic polymeric species from
aqueous solutions will be rapid in order to minimise
interfacial free energy.36 Earlier work using synthetic
polymers as skin mimics has demonstrated that, despite many obvious chemical and physical differences
to epidermal surfaces, carefully prepared artificial materials can display similar total surface energy values
compared to human skin.30,31 Thus an appropriate
skin substitute in terms of surface free energy should
exhibit similar adsorptive behavior to its natural counterpart. In turn, this should lead to the surface display
of the amphiphilic polymer irrespective of the substrate and a masking of the roughness and heterogeneity of the underlying material (epidermal cell or
synthetic polymer). We therefore reasoned that the
original substrate would be of less relevance at the
early stages of bacterial attachment than the presence
or absence of a densely packed layer of adsorbed copolymer.
The initial work involved a contact angle goniometry screen of model surfaces to act as physicochemical mimics for skin. Three different synthetic polymers—PDMS, PMMA, and PS—were chosen, adsorbed onto silanized glass slides via dip coating from
solution and/or spin coating, and contact angles were
measured on these substrates after droplet contact
times of 1 and 20 min. Three different liquids (water,
diiodomethane, and 1,2-ethanediol) were used to derive Lewis acid (␥+), Lewis base (␥−), Lifshitz–van der
Waals (␥LW), and total surface energy (␥tot) values according to the method of Good et al.37 The same methodology was used to derive surface energies of skin
from human volunteers, although contact angles for
these experiments were recorded after 1 min only. It
can be seen (Table I) that although PMMA displayed
the most similar value for ␥tot after 1 min, PS surfaces
were closest in terms of individual energy components to those of human skin samples. Although previous work has identified total surface energy as a key
factor in determining skin mimetic properties,30 we
wanted also to consider surface energy components,
particularly van der Waals and acid–base components, as skin contains varying amounts of proteins
and lipids that exhibit amphiphilic character. As these
components are likely to be crucial parameters in adhesion to synthetic substrates,38 as they determine
acid–base and induced dipole interactions at the surface, we adopted PS, which gave the closest overall
match of surface energy components, as the model
skin mimic and prepared smooth, good-quality PS
films for further experiments.
Adsorption of PEO-PPO copolymers has been previously reported by Bridgett et al.39and Amiji and
Park,40 who indicated that the bacterial fouling resistance of the resultant surfaces was the length of the
hydrophobic PPO block and its proportion in comparison to the hydrophilic PEO segments. This was
TABLE I
Surface Energies of Model Polymer Substrates and
Human Skin
Material
Time
(s)
PS
PS
PMMA
PMMA
PDMS
PDMS
Skin
60
1200
60
1200
60
1200
60
Surface Free Energy Components (mJ m−2)
␥sLW
␥s+
␥s−
␥sTotal
31.0
34.0
36.7
36.6
22.0
29.0
33.0
0.1
0.4
0.02
0.01
1.9
2.0
0.2
8.0
9.0
17.8
20.0
8.0
13.0
12.0
33.0
37.2
37.9
37.5
29.4
40.0
36.5
ADSORBED PEO-PPO COPOLYMERS ON SYNTHETIC SURFACES
ascribed to the higher affinities of longer hydrophobic
segments in the copolymer backbone with the hydrophobic base layer of poly(styrene) and the consequent
stronger anchoring of the PEO-PPO coating at the surface. We therefore chose a range of BASF Pluronic™
copolymers varying in block lengths and hydrophilic/
hydrophobic balances in order to obtain coatings that
most strongly bound to PS substrates. The surfaces
were initially characterized in terms of their advancing contact angles in water, and these varied from
extremely hydrophilic (␪w < 10°) to hydrophobic (␪w >
60°) in nature. Goniometry showed considerable dropto-drop variations in water contact angles across many
different PEO-PPO graft surfaces, suggesting uneven
coverage of the PS-silanized glass substrates by the
adsorbed copolymers. These surface heterogeneities
were similar to those recently observed by Green et
al.,41 who used AFM as part of experiments to probe
the structures of PEO-PPO coatings, and indeed
atomic force micrographs in our study showed the
presence of discrete domains of PEO-PPO overlying
partially the PS-silanized glass [Figs 2(a, b)]. However,
645
careful control of experimental conditions facilitated
the preparation of smooth homogeneous coatings,
with very low water contact angles (␪w < 12°), suggesting that adsorption of the copolymers had taken
place such that the PEO chains were extended into the
bulk solution. The highest quality coatings were obtained with Pluronic F127, although it was not possible to calculate the surface free energy components
of this amphiphilic copolymer via contact angle goniometry due to strong interactions with both polar and
nonpolar diagnostic liquids. AFM images obtained
under water indicated the presence of a surface layer,
which appeared smooth and largely featureless in topography, with surface roughness (Ra) values as low
as 20 Å in some cases. Few surface structures were
apparent, although at higher magnification polymer
aggregates were visible that were not seen in the underlying PS substrates. However, further AFM studies
in air of these adsorbed F127 surfaces revealed a degree of secondary structure after drying, with the appearance of pseudocrystalline domains. While some
areas of the substrates were smooth [Fig. 2(c)], the
Figure 2. Atomic force micrographs of copolymers on PS-silanized glass substrates. (a) PEO-PPO copolymer F68, x = y = 50
␮m. z = 58 nm. (b) PEO-PPO copolymer F127 showing surface heterogeneity, x = y = 50 ␮m, z = 28 nm. (c) PEO-PPO
copolymer F127 on PS-silanized glass prepared by optimized procedure, x = y = 20 ␮m, z = 13 nm. Smooth topology shown.
(d) Pseudocrystalline structure of PEO-PPO copolymer F127, x = y = 30 ␮m, z = 65 nm, with associated amorphous area
(left-hand side of surface) to show relative proportions. [Color figure can be viewed in the online issue, which is available at
www.interscience.wiley.com.]
646
MARSH ET AL.
majority of the surfaces exhibited the topography
shown in Figure 2(d) with the relative proportions of
the contributing structures clearly displayed.
Additional structural information on the adsorbed
PEO-PPO layers was obtained using ATR-FTIR spectroscopy. The F127-PS films were carefully peeled
from the silanized glass substrates and pressed against
a germanium ATR surface for spectra to be recorded.
The IR absorptions showed characteristic C−O
stretches at 1112 and 1098 cm−1 as expected from the
polyether backbone, but secondary analysis of peak
shapes indicated that the PEO-PPO copolymer was, as
suggested by AFM, present in at least partially crystalline domains. Comparison of IR spectra of PS-F127
with those of PEGs and oligo(ethyleneglycol) selfassembled monolayers (SAMs)42 showed a number of
similarities. For example, the bands in the PS-F127
spectra at 2890, 1345, 1283, 1149, and 963 cm−1 were
also present in spectra of crystalline PEG, which were
assigned to the CH2 symmetric stretch, the CH2 wag
(gauche conformation), the C−O and CH2 stretches,
and the CH2 twist and CH2 rocking modes of crystalline oligo(ethyleneglycol). These absorption bands
were not present in amorphous oligo(ethyleneglycol)
or in PPO spectra (Table II). The presence of these
pseudo-crystalline areas indicated a dense coverage of
the adsorbed PEO-PPO polymer at the surface, such
that the PEO blocks were able to order in a closepacked arrangement. Previous studies have indicated
that the formation of a dense “brush” structure at the
surface is a key factor in the resistance of grafted or
adsorbed hydrophilic polymers to bioadhesion,43 and
TABLE II
Infrared Absorption Modes of Adsorbed F127 Films and
Polymer Analogues
Polymer
Adsorption
Band (cm−1)
F127
Crystalline
PEG
Amorphous
PEG
2969
2890
2740
1467
2970
2930
2895
2950
2930 sh
2890 s
2885 s
2865 sh
2740
1470 m
1460 m
2865 b
2740 sh
2870
1460 m
1461 sh
1373
1342
1359
1344
1345 s
1280
1283 m
1241
1149
1112
1244 m
1149 s
1119 s
1102 vs
1062 m
963 s
1060
964
PPO
1352 m
1325 w
1296 m
1249 m
1140 sh
1107 s
1038 m
945 m
1296
1257
thus it seemed likely that our PS-F127 surfaces, as initially prepared, were ordered sufficiently such that
when hydrated a dense brush layer would be present
to suppress the adsorption of microorganisms.
We subsequently challenged the PS-F127 surfaces
with a clinically important marker organism, Serratia
marcescens, and the skin-borne pathogen, Staphylococcus epidermidis, and compared the numbers of attached
organisms with those adsorbed to cleaned glass slides,
silanized glass, and PS-coated glass under the same
conditions. Fluorescence micrographs of ethidium
bromide labeled Serratia marcescens [Fig. 3(a–d)]
showed a significant decrease in the number of organisms bound (or retained on rinsing) to the PS-F127
substrates relative to the control surfaces (PS, glass,
and silanized glass). The same pattern of bacterial adsorption was observed for S. epidermidis over a similar
number of experiments. Estimates of cell numbers at
the surface based on at least triplicate fields of view
indicated ∼3 × 106 CFU ⭈ cm−2 adsorbed to PS and
clean glass surfaces, ∼5 × 106 CFU ⭈ cm−2 to silanized
glass, and only 1–2 × 104 CFU ⭈ cm−2 attached to the
PS-F127 substrates. It was thus clear that the adsorbed
PEO-PPO copolymer coating effected a decrease in
bacterial attachment in vitro, and the results suggested
that even our simple dip-coating method, using appropriate amphiphilic copolymers, might provide an
effective barrier to cell adsorption in vivo.
However, the limitations inherent to conventional
microscopy, particularly the difficulty in detecting
and enumerating accurately very low numbers of attached bacteria across the required surface areas
(1cm2), and the complexity in chemically mapping the
surfaces by this technique, rendered further interpretation of the bacterial adsorption experiments problematic. For example, it was not possible via conventional microscopy alone to establish whether the
Pluronic copolymer film remained unchanged on the
surface throughout the assay, had retained its tightpacking arrangement, or had simply been removed
during immersion in the bacterial culture suspensions,
in effect acting as a sacrificial layer to prevent cell
adsorption. In addition, the presence or extent of any
conditioning layer overlying the F127-PS test surfaces
could not be determined. As the adsorption of biopolymers is considered to be the first stage in bacterial
attachment and subsequent colonization, it was important to ascertain whether the outermost layer of
our model substrates was compromised by bacterial
challenge.
We therefore carried out further spectroscopic and
microscopic analysis to compare the structures of the
test surfaces before and after incubation in bacterial
cultures. A dye-labeled derivative of F127 PEO-PPO
copolymer was prepared by reaction with FITC and
adsorbed to the model PS surfaces as before. Fluorescence spectra recorded through glass coverslips with
ADSORBED PEO-PPO COPOLYMERS ON SYNTHETIC SURFACES
647
Figure 3. Fluorescence micrographs of Serratia marcescens stained with ethidium bromide on surfaces. (a) PS-silanized glass,
(b) clean glass, (c) silanized glass, and (d) F127-PS-silanized glass. [Color figure can be viewed in the online issue, which is
available at www.interscience.wiley.com.]
adsorbed FITC-F127 before incubation in the bacterial
suspensions displayed the expected characteristic
emission band centered at 510 nm following excitation
at 495 nm, and this mode was retained after immersion in bacterial cultures for 24 h although diminished
in intensity by ∼50%. While the low amounts of the
labeled polymer on the coverslips rendered quantitation difficult, these spectra confirmed that the PEOPPO copolymer coating remained adsorbed to the PS
substrate during bacterial challenge but indicated a
reduction in film thickness. FTIR spectra were recorded for F127-PS surfaces after bacterial adsorption
assays and compared with those obtained for freshly
prepared PS-F127 (Fig. 4). As can be seen in the spectra
from the incubated samples, the F127 C−O stretch at
1112 cm−1 is partially obscured by a broad band centered at 1098 cm−1. Although a shoulder absorption
band was visible at 1110 cm−1, the data suggested that
the surface was partially covered by adsorbed biopolymers, most likely excreted by the bacteria during the
incubation period. In addition, amide I and II bands
were also detected at 1641 cm−1, due to the adsorption
of proteins in the nutrient broth used to culture the
organisms. The spectra clearly showed PS C−C bands
at 1601 cm−1 again indicating, as suggested by the
fluorescence spectra, that the F127 polymer layer was
retained but that the thickness of the copolymer coating was reduced. As a consequence of the lack of unambiguous characteristic C−O bands in these spectra
and the difficulties in obtaining an accurate assessment of PEO-PPO removal, an XPS study was then
carried out on the surfaces. Spectra were recorded for
the following samples: F127 coated onto PS on silanized glass (S1); PS-F127 after incubation in phosphate-buffered saline (PBS) for 24 h (S2); PS-F127 after
incubation with S. epidermidis in PBS for 24 h (S3); and
PS-F127 after incubation with S. marcescens in PBS for
24 h (S4). The spectra obtained were curve fitted,29
normalized for relative atomic percentages present in
each sample; the data of relative atomic percentages
from each of the four surfaces are shown in Table III.
The C1s peak in the PS-F127 spectrum arising from
electrons in C−C and C−H bonds served as a “calibration.” The total area under this peak (AC1S) could be
related to the respective mole fractions (␹) of the constituents of the sampled surface layers according to
expression (1) below, as these signals could only arise
from coatings on top of the glass slides.
ACIS(C–C, C-H) = ␹PS + ␹F127 + ␹PDMS
(1)
Similarly, the second C1s peak, with energies corresponding to C−O bonds, could be attributed to the
648
MARSH ET AL.
Figure 4. ATR-FTIR spectra of polymer surfaces. (a) F127PS and (b) F127-PS after incubation with bacterial culture for
24 h.
C−O groups in F127 (␹F127), thus giving a linear relation between peak area and F127 content:
ACls 共C–O兲 = ␹F127
(2)
In order to interpret these results in terms of surface
compositions and layer thicknesses, it was assumed
that the both the PS and PEO-PPO coatings adsorbed
in a smooth, homogeneous manner, and also that the
C−O peak arose only from ether linkages in F127. As
supplied, F127 contains 31% PPO and 69% PEO, giving a “constitutive formula weight” of the polymer of
32 g mol−1, with one oxygen atom per repeat unit (i.e.,
TABLE III
XPS Assignments and Elemental Compositions of
Polymer Films
Binding
Energy
(eV)
Assignment
S1
S2
S3
S4
285.00
286.60
292.20
532.85
102.10
104.05
133.15
400.50
C1s (C—C, C—M)
C1s (C—O)
C1s (Shake-up)
O1s
Si2p (1)
Si2p (2)
P2p
N1s
41.2
21.9
61.9
9.7
1.3
18.1
8.4
Trace
Trace
62.8
5.4
1.3
19.4
10.6
Trace
51.6
8.0
Trace
24.2
13.6
1.5
Trace
Trace
a
Relative Atomic Percentagea
25.5
11.4
Trace
S1: PS-F127 as prepared. S2: PS-F127 after incubation in
phosphate buffer saline (PBS) for 24 h. S3: PS-F127 after
incubation with S. epidermidis in PBS for 24 h. S4: PS-F127
after incubation with S. marcescens in PBS for 24 h.
50% of the mass of each “monomer” unit). Thus for
surface S1 (PS-F127 prior to immersion), assuming
that the C1sC−O peak arose only from F127, then
the contribution of the PEO-PPO copolymer to
the C1sC−C, C−H peak area would be expected to
be twice that of the C1sC−O peak area. The experimentally determined relative peak areas for
C1sC−O and C1sC−C, C−H were 21.9 and 42.1% respectively, suggesting that almost all of the C1sC−C, C−H
peak observed in S1 was derived from absorption in
the F127 layer. In addition, the O1s peak area found in
spectrum S1 corresponded to a relative atomic percentage of 25.5%, of which 21.9% could be attributed
to the C−O oxygen atoms in the PEO-PPO copolymer.
This is because an XPS spectrum of F127 alone would
be expected to give equivalent peak areas for O1s and
C1sC−O absorptions. The remaining O1s absorptions
[i.e., the difference between the peak areas observed
for O1s and C1s(C−O) − 3.6%] were most likely to
have been due to oxygen in the underlying siloxane
layers. As the penetration depth of XPS in these experiments was 1–10 nm, dependent on incident angle,
the relative C1sC−C, C−H:C1sC−O:O1s ratios (∼2:1:1) suggested the beam was sampling primarily the F127
layer. The small Si2p [1] peak (11.4%) observed in
these spectra due to SiO in the underlying glass or
PDMS indicated that the PS and F127 layers were in
places thin or patchy.
XPS spectra of PS-F127 surfaces after immersion in
buffer (S2) showed a substantial increase in the
C1sC−C, C−H:C1sC−O ratio, with an extra peak arising
from C1sshake-up photoelectrons. As these peaks, resulting from absorptions in conjugated C=C bonds, were
not observed in S1, it was apparent that immersion in
PBS for 24 h removed some of the PEO-PPO layer to
expose the PS below. Calculation of the relative peak
areas for C1s C−O (F127 copolymer, 9.7%) and
C1sC−C, C−H (PS, F127 19.4%) showed that 42.5% of the
C1sC−C, C−H peak area was due to PS, indicating an
approximate 50% reduction in the thickness of the
PEO-PPO layer. This partial removal of the F127 coating by immersion in buffer was also demonstrated by
a decrease in the ratio of C1sC−O to O1s peak areas,
suggesting that an increased proportion of the glass
and silanized surface was being sampled. However, in
S2 the peak area due to Si absorptions was lower than
that found in S1, which indicated some surface inhomogeneity, and the presence of a small phosphorus
peak showed that some phosphate buffer was adsorbed onto the surfaces. The XPS spectrum for surface S3 (PS-F127 after incubation in Staphylococcus epidermidis) was essentially the same as that obtained for
S2, except that the phosphorus peak was not observable, and a small peak attributable to nitrogen was
present. In this sample it was not possible to state
unambiguously the origin of the C1sC−O peaks (previously arising only from F127 C−O bonds), as these
ADSORBED PEO-PPO COPOLYMERS ON SYNTHETIC SURFACES
may also have been present from C−O groups in exopolysaccharides (EPS) excreted from the bacteria. The
appearance of nitrogen signals provided additional
evidence that bacterial exudates were adsorbed to the
surfaces, albeit in trace amounts. For sample S4 (PSF127 after incubation in Serratia marcescens) the nitrogen and phosphorus peaks were again present, and
the C1sC−C, C−H absorption was correspondingly decreased. The known tendency of bacterial species, especially staphylococci, to excrete significant quantities
of EPS, combined with the appearance of trace N and
P signals, strongly suggests that the surfaces in these
samples, despite the presence of a PEO-PPO layer, did
at least display partial coverage of adsorbed biopolymers even if present in very small amounts.
In order to obtain further information about the nature of the polymer surfaces and any adsorbed biopolymers and cells, AFM experiments were performed on
the surfaces following incubation with the bacterial
and rinsing. The micrographs [Fig. 5(a–d)] were in
agreement with the previous fluorescence microscopy
studies, in that a high number of attached microorganisms were observed on model PS and glass surfaces:
no organisms were detected on PS-F127 samples. In
addition, fibrillar structures were observed on PS surfaces [Fig. 5(a,b)] after incubation in Serratia marcescens
649
and S. epidermidis cultures, respectively, which, on the
basis of the prior FTIR and XPS studies, were most
likely to have been bacterial EPS aggregates. The lack
of similar features on the PS-F127 films [Fig. (5c,d)]
suggested that although FTIR and XPS implied the
presence of EPS on these surfaces, the extent of biopolymer adsorption was substantially smaller. Pseudocrystalline domains, previously observed on PSF127 surfaces, were not apparent on these films after
bacterial incubation experiments due possibly to the
partial desorption of the PEO-PPO copolymer, as indicated by fluorescence spectroscopy and XPS. This
lack of surface order may also have been a result of
molecular level interactions with small amounts of
bacterial exudates and/or nutrient buffer components.
Nevertheless, the absence of microorganisms on the
F127-PS test surfaces strongly suggested that the adsorbed copolymer films were still effective as antibacterial attachment coatings after 24 h.
However, the true extent of bacterial adsorption to
the PEO-PPO surfaces could not be ascertained from
spectroscopic studies or microscopy, owing to the inherent experimental difficulties and possible subjectivity in detecting low numbers of cells by optical
methods. In order to increase the sensitivity of the
adsorption assay experiments, bacterial cultures of S.
Figure 5. Atomic force micrographs of surfaces after incubation with bacterial cultures for 24 h. (a) PS-silanized glass slide,
x = y = 20 ␮m, z = 934 nm; incubation in S. marcescens. (b) PS-silanized glass slide, x = y = 23.4 ␮m, z = 610 nm; incubation
in S. epidermidis. (c) F127-PS-silanized glass slide, x = y = 10 ␮m, z = 143 nm; incubation in S. marcescens. (d) F127 PS-silanized
glass slide, x = y = 25 ␮m, z = 3.18 nm; incubation in S. epidermidis. [Color figure can be viewed in the online issue, which is
available at www.interscience.wiley.com.]
650
marcescens and S. epidermidis were labeled with 14C
glucose prior to incubation with test surfaces (PS-F127
surfaces on glass, and with control PS, silanized glass,
and clean glass substrates), which enabled consistent
detection down to 102–103 organisms per sample.
Figure 6(a) shows the results for adsorption of S.
marcescens to chromic acid-cleaned glass, silanized
glass, PS-coated glass, and PS-F127 coated glass. In
accord with previous AFM and fluorescence microscopy observations, an average decrease of one order of
magnitude in the attachment of S. marcescens to PSF127 samples was observed, compared to PS controls.
The overall number of cells attached to PS-F127 surfaces (average 2.8 × 104 ± 1.44 × 103 organisms per
polymer film) was significantly lower (n = 6, P =
0.0002) than that on PS control slides (2.0 × 106 ± 2.2 ×
104 organisms) and on silanized glass (1.4 × 105 ± 1.0
× 105 cells). These data are in agreement with many
previous observations and theoretical derivations in
that bacterial attachment was seen to be generally
higher to hydrophobic substrates in aqueous environments.44 The high standard deviation of the latter
samples may have been a result of the efficiency of
silanization of the previously cleaned glass slides, as
attachment of S. marcescens to cleaned glass slides (6.0
× 104 ± 2.3 × 102 organisms) was relatively low, reflecting the less favorable interfacial energy of adsorption to hydrophilic substrates.
In order to evaluate the performance of the F127coated “skin model” PS surfaces, a second set of adsorption experiments was carried out, with both S.
marcescens and S. epidermidis. The results [Fig. 6(b)]
were in agreement with the previous set, showing a
significant difference (n = 6, P = 0.0057) between the
numbers of S. marcescens attached to PS-F127 samples
(7.0 × 104 ± 5.1 × 104 cells per polymer film) and PS
MARSH ET AL.
control surfaces (5.0 × 105 ± 1.1 × 105 organisms). Similarly, the number of S. epidermidis retained on PS-F127
(3.6 × 104 ± 1.1 × 104 organisms) was one order of
magnitude smaller (n = 6, P = 0.0005) than that adhered to PS model surfaces (2.9 × 105 ± 4.1 ×104 organisms). The variation in the magnitude of attachment observed for the two bacterial strains can most
probably be explained in terms of the different cell
concentrations used in the individual assays (108
CFU ⭈ mL−1 for S. marcescens, and 107 CFU ⭈ mL−1 for
S. epidermidis): however, the relative degrees of adhesion to the PS-F127 and PS surfaces were seen to be
consistent for both types of microorganism. The radiolabeling experiments thus provided further confirmation that the adsorbed PEO-PPO coatings were effective in reducing cell attachment to PS model skin surfaces in vitro.
CONCLUSIONS
The results from this study show that an experimentally facile dip-coating method can be used to apply
commercially available PEO-PPO block copolymers to
synthetic substrates designed to act as physicochemical models for human skin. Furthermore, spectroscopic and microscopic analysis indicates that even
using this simple adsorption protocol highly ordered
and densely packed layers of PEO-PPO copolymer
were deposited on the model surfaces. Fluorescence
and atomic force microscopies established that adsorption of two bacteria implicated in skin infection,
Staphylococcus epidermidis and Serratia marcescens, was
suppressed significantly on the PEO-PPO-treated surfaces relative to PS substrates over a 24 h time period.
Figure 6. Attachment of bacteria to surfaces after 24 h. (a) S. marcescens to F127-PS, PS, clean glass, and silanized glass. (b)
S. marcescens and S. epidermidis to F127-PS and PS. Error bars represent standard deviations. ANOVA single factor analysis
was used to derive p values (n = 6). Significant differences were observed for S. marcescens attached to F127-PS compared to
PS in both experiments (p = 0.00019, 0.0057), and for S. epidermidis to F127-PS compared to PS (p = 0.000528).
ADSORBED PEO-PPO COPOLYMERS ON SYNTHETIC SURFACES
Radiolabeling experiments confirmed a reduction in
bacterial attachment to adsorbed PEO-PPO of at least
one order of magnitude compared to PS model surfaces. The results suggest that practical applications
for these polymers as antimicrobial or surface protectants should follow if the desired copolymer structures can be produced as densely packed ordered domains on biological substrates.
We thank Reckitt Benckiser Healthcare and BBSRC for
financial support (BBSRC Industrial CASE), Simon Young
and Vanessa Peters, University of Portsmouth for help with
contact angle goniometry and microbiological studies, and
Drs. Peter Eaton and Maureen Stone, University of Portsmouth, for helpful discussions. We also thank one of the
referees for helpful comments.
16.
17.
18.
19.
20.
21.
22.
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