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Applied and Environmental Microbiology, November 1999, p. 4995-5002, Vol. 65, No. 11
Macromolecular Science Department, Institute
of Food Research, Reading Laboratory, Reading RG6 6BZ, United
Kingdom
Received 11 May 1999/Accepted 17 August 1999
A systematic investigation into the effect of surface chemistry on
bacterial adhesion was carried out. In particular, a number of
physicochemical factors important in defining the surface at the
molecular level were assessed for their effect on the adhesion of
Listeria monocytogenes, Salmonella typhimurium,
Staphylococcus aureus, and Escherichia coli.
The primary experiments involved the grafting of groups varying in
hydrophilicity, hydrophobicity, chain length, and chemical
functionality onto glass substrates such that the surfaces were
homogeneous and densely packed with functional groups. All of the
surfaces were found to be chemically well defined, and their measured
surface energies varied from 15 to 41 mJ · m The prevention of contamination
caused by pathogenic microorganisms during the manufacture, processing,
and packaging of food is of considerable importance to public health
and consequently is a major issue for industry (30). In
particular, there is increasing concern associated with the
contamination arising from bacterial biofilms which develop on
materials used during food manufacture (6, 19, 40). These
communities of bacteria, often embedded in a matrix of organic polymers
exuded by the cells, can be extremely difficult to remove, and complete
eradication of the pathogens is difficult, time-consuming, and
expensive (27). In general, the formation of bacterial
biofilms is believed to take place over at least three stages: a
reversible adsorption step (26), primary adhesion of
microorganisms to a surface, and colonization (9). The rates
of these processes vary widely depending on the environmental
conditions and the type of microorganisms, but the adhesion and
colonization stages are considered to be relatively slow compared to
the first step of cell adsorption (17). In principle, it
should be possible to retard, if not prevent, the formation of biofilms
on substrates by using materials to which bacteria cannot initially
attach, and such a material or surface coating would be of considerable
commercial interest (7). In practice, however, synthetic
materials that are capable of preventing bacterial adsorption have
proved rather elusive, despite a significant volume of research
(8, 11). Properties of the substrate, such as hydrophobicity
(32), hydrophilicity (23), steric hindrance
(24), roughness (22), and the existence of a
"conditioning layer" at the surface (1), are all thought to be important in the initial cell attachment process.
In addition, there are large discrepancies in the data available on the
adhesion of microorganisms to various synthetic materials. These
disparities are mainly due to the different experimental conditions and
protocols used in different laboratories as well as to the fact that
the substrates employed are sometimes incompletely characterized both
in terms of their functionality, i.e., the actual chemistry of the
groups present, and in terms of their density at the surface.
Consequently, it is often difficult to draw a meaningful comparison
between results reported by different authors, and no simple
structure-function correlation has yet emerged (33). The
objective of this investigation was to carry out a comprehensive
comparative study of bacterial adsorption by using a broad range of
surfaces with controllable and precisely defined chemistry, such that
various chemical and physicochemical factors which may be important in
defining the surface at the molecular level could be assessed for their
effect on the adhesion of the representative food pathogens
Listeria monocytogenes, Salmonella typhimurium,
Staphylococcus aureus, and Escherichia coli.
Chemicals.
Poly(ethyleneoxide)-monomethylether (MeO-PEO)
(Mn = 350, 500, 750, 2,000, and 5,000),
2-[2-(2-methoxy)ethoxy]acetic acid (MeO-PEO-3-CO2H), 3-mercaptopropanoic acid, 2-aminoethanethiol,
1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride
(EDC), 4,4'-azobis(4-cyanovaleric acid) (ACVA), 3-aminopropyltriethoxysilane (APTES), ethanoyl chloride, butanoyl chloride, octanoyl chloride, 2-ethylhexanoyl chloride, decanoyl chloride, oleoyl chloride, myristoyl chloride, phenolphthalein (A.C.S.
reagent), hydrogen peroxide, triethylamine, toluene,
1,1,2-trichlorotrifluoroethane (A.C.S. reagent), tetrahydrofuran (THF),
dansyl chloride, and sodium carbonate were purchased from Aldrich
Chemical Company (Gillingham, United Kingdom). Perfluorobutanoyl
chloride and perfluorooctanoyl chloride were purchased from Lancaster
Synthesis (Lancaster, United Kingdom). Methanol (high-pressure liquid
chromatography [HPLC] grade), 2-propanol (HPLC grade), acetone (AR
grade), diethyl ether (HPLC grade, peroxide-free), sodium hydrogen
carbonate, sodium hydroxide, potassium dichromate, sulfuric acid, and
p-toluene sulfonoyl chloride were purchased from Fisher
Scientific (Loughborough, United Kingdom). Azobis(isobutyronitrile)
(AIBN) was purchased from Acros Organics (Loughborough, United Kingdom)
and recrystallized from methanol prior to use. 2-Mercaptoethanol,
ethanol (AR grade), and chloroform (AR grade) were purchased from BDH
(Poole, United Kingdom). Sulfo-SDTB was purchased from Pierce
(Rockford, Ill.). N-Methylacrylamide was purchased from
Monomer-Polymer Laboratories (Trevose, Pa.). Krytox acid terminal
perfluoropolyether was obtained from DuPont (Deepwater, N.J.).
Cytochrome c (from bovine heart) and bovine serum albumin
(BSA) were obtained from Sigma (Poole, United Kingdom).
D-[U-14C]glucose and 3H-acetic
anhydride were obtained from Amersham Life Sciences (Amersham, United
Kingdom). Unless otherwise stated, all chemicals were used as received.
Growth and maintenance of microorganisms.
Bacterial cultures
were maintained on Tryptone Soya Agar (Oxoid) at 4°C. Stock cultures
were grown statically overnight at 37°C in coryneform broth
(containing [in grams per liter] tryptone, 10; yeast extract, 5;
sodium chloride, 5; and glucose, 5; pH 7.2) for L. monocytogenes C200 type 2 and in yeast-dextrose broth (containing [in grams per liter] peptone, 10; beef extract, 8, sodium chloride, 5; glucose, 5; and yeast extract, 3; pH 6.8) for S. typhimurium LT2 50185, S. aureus NCDO 949, and
E. coli NCFB 1989. Aliquots (100 ml) of stock culture
in sterile Eppendorf tubes were drop-frozen in liquid nitrogen and
stored at Preparation and characterization of silylated glass
surfaces.
Glass coverslips (13-mm diameter; Chance Propper, Ltd.,
Smethwick, United Kingdom) were hydrolyzed by immersion in sodium hydroxide (aqueous, 5 M) for 1 h and washed thoroughly with
deionized water. They were then soaked in fresh piranha solution (70%
sulfuric acid, 30% hydrogen peroxide) for 1 h, washed with water,
and dried in air. The glass surfaces were prepared immediately prior to silylation with APTES by drying them in a hot oven for 30 min. The
coverslips were then immersed in a solution containing 95% acetic acid
(1 mM) in methanol, 4% water, and 1% APTES for 30 min at room
temperature with occasional gentle shaking. The resultant amine-functional surfaces were finally washed with methanol (three times with 50 ml) and cured in a hot oven for 30 min.
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Bacterial Adhesion at Synthetic Surfaces
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
2.
Protein adsorption experiments were performed with
3H-labelled bovine serum albumin and cytochrome
c prior to bacterial attachment studies. Hydrophilic
uncharged surfaces showed the greatest resistance to protein
adsorption; however, our studies also showed that the effectiveness of
poly(ethyleneoxide) (PEO) polymers was not simply a result of its
hydrophilicity and molecular weight alone. The adsorption of the two
proteins approximately correlated with short-term cell adhesion, and
bacterial attachment for L. monocytogenes and E. coli also correlated with the chemistry of the underlying
substrate. However, for S. aureus and S. typhimurium a different pattern of attachment occurred,
suggesting a dissimilar mechanism of cell attachment, although
high-molecular-weight PEO was still the least-cell-adsorbing surface.
The implications of this for in vivo attachment of cells suggest that
hydrophilic passivating groups may be the best method for preventing
cell adsorption to synthetic substrates provided they can be grafted uniformly and in sufficient density at the surface.
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
70°C prior to use. Prior to the first adhesion
experiments, growth curves were established for the bacteria by
correlating plate counts and optical density measurements. Bacteria
plated from the stationary phase showed no significant loss in
viability after 24 h, as determined by plate counts. Cell
viability after the adsorption experiments was qualitatively confirmed
by using confocal microscopy, which clearly showed the formation of
bacterial colonies on the grafted surfaces and plate counts, by using
polymer beads rather than films but with the same surface functionality.
1). The reaction was carried out in the
dark at room temperature for 2 h. The coverslips were then washed
twice with chloroform (5 ml) and three times with ethanol (5 ml) and
dried in air, and the fluorescence spectra were recorded. For UV
spectroscopy, three coverslips were placed in a quartz cuvette to
enable sufficient peak absorptions to be recorded.
Synthesis of reagents. (i) MeO-PEO-CO2H. Synthesis was carried out as described previously (13). The protocol described below is for MeO-PEO-5000, but a similar procedure was used for other PEO oligomers. High-molecular-weight MeO-PEO-OH (5.0 g, 1 mmol) was dissolved in anhydrous DMF (100 ml) with 1.1 eQ of succinic anhydride (0.1 g) in a dry, three-necked 250-ml round-bottomed flask and heated to 100°C under an atmosphere of dry nitrogen overnight. Upon cooling the mixture was concentrated under reduced pressure, and the polymer was purified by precipitation into cold ether (three times, 250 ml), followed by drying in a vacuum oven at 40°C overnight.
(ii) MeO-PEO acyl chlorides. High-molecular-weight carboxyl-terminated MeO-PEO (5.1 g, 1 mmol) was dissolved in dry toluene (100 ml) in a 250-ml three-necked round-bottomed flask. Oxalyl chloride (0.375 g, 3 mmol) was added, and the mixture was brought to reflux under an atmosphere of dry nitrogen and stirred at reflux overnight. Upon cooling the solvent and excess oxalyl chloride were removed under reduced pressure, and the resultant MeO-PEO-acyl chloride was used immediately. The acyl chloride of 2-[2-(2-methoxy)ethoxy]acetic acid (MeO-PEO-3-COCl) was synthesized in the same way.
(iii) Krytox acyl chloride. Krytox (5.0 g) was dissolved in 1,1,2-trichlorotrifluoroethane (100 ml) in a 250-ml three-necked flask. Oxalyl chloride (3 ml) was added, and the mixture was brought to reflux under an atmosphere of dry nitrogen and stirred at reflux overnight. Upon cooling the solvent and excess oxalyl chloride were removed under reduced pressure, and the resultant acid chloride was used immediately.
(iv) Poly(N-methylacrylamide) polymers. In a thick-walled tube, monomer (N-methylacrylamide, 10 g) was dissolved in 2-propanol (40 ml) with chain transfer agent (0.344 mmol) and initiator (AIBN or ACVA [see below], 2.83 mmol). This mixture was degassed by freeze-thaw cycles under vacuum at least three times. The tubes were then placed in a thermostatted oil bath at 65°C for 24 h. After it cooled to room temperature, the mixture was concentrated under reduced pressure, and the residue was added to diethyl ether (250 ml) to precipitate the polymer. This was filtered and the residue was redissolved in THF and reprecipitated into diethyl ether (three times, 250 ml) to leave the purified polymer as a colorless precipitate which was then dried in vacuo at 20°C overnight.
The molecular weight and functionality of the poly(N-methylacrylamide) polymers were controlled with the use of functional initiators and suitable chain transfer agents. Carboxyl-terminated polymers were obtained with the use of 3-mercaptopropanoic acid as a chain transfer agent and ACVA as the free radical initiator. The molecular weight of carboxyl-terminated poly(N-methylacrylamide) (~100 mg) was determined by titration of dissolved polymer in deionized water (50 ml) with freshly prepared sodium hydroxide solution (10 mM). The endpoint was either determined potentiometrically or by using phenolphthalein solution as an indicator. Amine- and hydroxy-terminated polymers were obtained by using AIBN as the free radical initiator and 2-aminoethanethiol or 2-mercaptoethanol as the chain transfer agent.Chemical modification of surfaces. (i) Treatment with carboxyl-terminated poly(N-methylacrylamide). Silylated coverslips were placed in dilute hydrochloric acid (pH 6) and cooled to 0°C. Carboxyl-terminated poly(N-methylacrylamide) (1.0 g) was added to the buffer solution, followed by EDC (three times, 200 mg) every 30 min with gentle stirring. After the addition was complete, the surfaces were left at 0°C for a further 72 h. The surfaces were then rinsed repeatedly with deionized water (five times, 250 ml) and dried in air.
(ii) Treatment with acid chlorides. Glass coverslips were placed in dry chloroform (40 ml) and triethylamine (10 ml); to this the required acid chloride (5 ml) was added, and the mixture was left at room temperature under nitrogen with gentle stirring for 2 h. The coverslips were then washed with chloroform (four times, 50 ml) and dried in air.
(iii) Treatment with Krytox acid chloride. Glass coverslips were placed in 1,1,2-trichlorotrifluoroethane (10 ml) and triethylamine (3 ml) containing Krytox acid chloride (3 g) and left at room temperature with gentle stirring for 1 h; the coverslips were then washed in a Soxhlet apparatus with 1,1,2-trichlorotrifluoroethane overnight and dried in air.
To remove possible contamination remaining after grafting, the substrates were cleaned by sequential washing with ethanol (three times, 100 ml) and distilled water (three times, 100 ml), followed by rinsing with water and drying in dust-free air.Assessment of bacterial attachment to synthetic surfaces.
To
carry out radioactive labelling of microorganisms, modified coryneform
broth (6 ml, containing a growth-limiting concentration of glucose [1
g · liter1]) for cultures of L. monocytogenes or modified yeast-dextrose broth (6 ml,
containing glucose [1 g · liter1]) for
cultures of S. typhimurium, S. aureus, or
E. coli were inoculated with 50 µl of thawed stock
culture. Aliquots of D-[U-14C]glucose were
added to a final concentration of 20 µCi · ml1
from a 1-mCi stock solution (230 to 370 mCi · ml1;
a sterilized, aqueous solution containing 3% ethanol). The cultures were incubated, statically, at 37°C for 24 h.
Determination of absolute cell counts. Aliquots (200 µl) of unlabelled bacterial culture were enumerated via a serial dilution method with Tryptone Soya Agar and incubation at 37°C for 24 h. The numbers of viable cells determined this way were compared with the scintillation counts from equivalent aliquots (200 µl) of radiolabelled bacteria. In addition, the scintillation readings were correlated with total bacterial numbers obtained via optical density measurements and microscopy.
Adsorption of proteins to synthetic surfaces.
The method to
prepare radioactively labelled proteins was adapted from that used by
Freeman and Parish to label heparan (12). Cytochrome
c or BSA (20 mg in each case) was dissolved in
NaHCO3 (aqueous 0.5 M, 1 ml) containing 10% (vol/vol)
methanol in a sealed 15-ml reaction vial and cooled to 0°C in an ice
bath. 3H-acetic anhydride (0.1 ml, 10 mCi; 500 mCi · mmol1) in toluene was added, and the mixture was stirred
for 3 h at 0°C. The mixture was acidified to pH 7.0 with acetic
acid and allowed to warm to room temperature, and the toluene was
removed under a stream of nitrogen. The solution was desalted by using a PD-10 column (Pharmacia Biotech, St. Albans, United Kingdom) equilibrated and developed with aqueous ethanol (10% [vol/vol]). Column fractions containing radioactive material appearing ahead of the
3H-acetate peak were pooled, sodium azide was added (0.1%
[wt/vol]), and the solution was finally stored at 4°C. Protein
concentration was determined by using the Lowry method for BSA and from
a spectrophotometric calibration curve (408 nm) for cytochrome
c.
Pretreatment of substrates with BSA and cell exudate. Native BSA (37.5 µg) was added to MOPS (50 mM, pH 7.0, 10 ml) in 15-ml screw-capped bottles. Glass coverslips (13-mm in diameter), previously stored in absolute ethanol, were transferred to the protein solutions. The bottles were incubated, with gentle shaking, for 1 h (unless otherwise stated) at 37°C. The coverslips were then rinsed in sterile MOPS (50 mM, pH 7.0, 10 ml) and transferred to 15-ml screw-capped bottles containing sterile MOPS (50 mM, pH 7.0, 10 ml). Aliquots (200 µl) of 14C-labelled L. monocytogenes suspension were added, and the bottles were incubated, with gentle shaking, for 24 h as described above. Glass coverslips were subsequently rinsed in sterile MOPS (50 mM, pH 7.0, 10 ml) and transferred to scintillation vials for counting by the standard method.
To obtain cell exudate, modified coryneform broth (6 ml, containing a growth-limiting concentration of glucose [1 g · liter1]) was inoculated with 50 µl of thawed stock
culture and subsequently incubated, statically, at 37°C for 24 h. The culture (6 ml) was then transferred to Eppendorf tubes and
centrifuged (3 min, 13,000 rpm). Aliquots (200 µl) of the resulting
supernatant were then transferred to sterile MOPS (10 ml) in 15-ml
capped bottles, and glass coverslips were aseptically transferred to
the bacterial exudate solution. The bottles were shaken at constant
temperature for 1 h before the glass coverslips were rinsed in
sterile MOPS (50 mM, pH 7.0, 10 ml) and transferred to sterile MOPS (10 ml) in 15-ml capped bottles. Radiolabelled BSA (37.5 µg) was then added, and protein adsorption was allowed to proceed for 1 h at 37°C. The coverslips were subsequently rinsed twice in MOPS (50 mM,
pH 7.0, 10 ml) and transferred to scintillation vials for counting as
described above.
All data from protein and cell adsorption assays were averaged over at
least 10 replications, and standard deviations from the mean were
calculated. The results obtained (see Fig. 2 to 5) are averages of
these replications ± the standard errors.
Other analytical methods. Fourier transform-attenuated total internal reflection infrared spectra were recorded on a Perkin-Elmer 1600 spectrometer (Perkin-Elmer, Seer Green, United Kingdom) with a SpectraTech baseline attenuated, total internal reflectance apparatus by using a 45° germanium flat-face prism purchased from Nicolet Instruments (Nicolet, Warwick, United Kingdom). Fluorescence spectra were obtained by using a Perkin-Elmer LS50B Luminescence Spectrometer equipped with a front-face sample cell. Nuclear magnetic resonance (NMR) spectra were recorded on a Jeol-EX 270 NMR spectrometer (Jeol-UK, Welwyn Garden City, United Kingdom) by using CDCl3 or CD3SOCD3 as the solvents, with residual proton signals as the internal reference. UV spectra of modified surfaces were recorded on a Perkin-Elmer Lambda 15 spectrometer.
Contact angle measurements were carried out on Krüss G-10 goniometer (Krüss GmbH, Hamburg, Germany) with a G-211 environmental cell and fitted with square pixel video capture camera; analysis of captured images was done by using Krüss Drop Shape Analysis software. Glass microscope slides derivatized as described above were used. The slides were placed in a Krüss 211 environmental chamber, and advancing and receding contact angles were measured by using three diagnostic liquids: diiodomethane, water, and ethylene glycol. Surface free energy values and the relative contributions of Lifshitz-van der Waals, electron donor, and electron acceptor components were calculated by using the method of van Oss et al. (38).| |
RESULTS |
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Preparation and characterization of surfaces. Modified silica glass, in the form of 13-mm microscope coverslips, was chosen as a standard substrate for adhesion studies because of convenience of handling. As illustrated in Fig. 1, the coverslips were treated with APTES via the method of Durfor et al. (10) to give a slightly hydrophobic amine-functional surface, to which pendant groups of various degrees of chemical functionality, hydrophobicity, hydrophilicity, size, and hydrogen-bonding ability were grafted. Further derivatization reactions of APTES-modified coverslips were carried out by coupling acid chlorides or carboxylic acids to amine groups to generate short- and long-chain hydrophobic, hydrophilic, fluorocarbon, alkylamide, and alkylether surfaces. The functionalities of the surfaces grafted onto these SiO2-APTES substrates are shown in Table 1.
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9
mol · cm2 or less. However, fluorescence
spectroscopy proved insufficiently sensitive for monitoring the extent
of further surface grafting to the 13-mm coverslips directly, and
silica beads (100 µm in diameter) were therefore used as a
higher-surface-area model to assess the extent of modification.
Titration of amine groups on these beads by a variant of the method of
Guar and Gupta (16) gave, within experimental error,
essentially the same result, with a density of amine groups of 5 × 1010 mol · cm2 (8.9 × 108 mol · g1 [corresponding to ca.
1 amine group per 36 Å2]). This number was reduced to
5 × 1011 to 7 × 1011 mol
· cm2 after the grafting reactions carried out with the
EDC coupling procedure, indicating that at least 87% of the surface
amine groups were successfully derivatized. For grafting via acyl
chlorides, titrations showed that more than 99% of the available amine
groups were derivatized under the reaction conditions used.
Second, the free energies of the derivatized surfaces were determined
by dynamic contact angle measurements, and the results obtained (Table
2) were in good agreement with values
reported for the same or similar functional groups (14).
These data suggest that, under the conditions used, APTES substrates
were uniformly derivatized via our grafting chemistry with virtually no
underlying amine or silica surface exposed. Atomic force microscopy
(AFM) also provided evidence that the surfaces were smooth and
physically homogeneous to a submicron level (not shown).
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), S-C6 (
), and hydrophobized SiO2
(
). On the right, the adsorption of BSA against concentration is
shown: S-NH2 (
) and S-C8F (2) than that of BSA (3.3 pmol · cm2). However, adsorption of cytochrome c to
hydrophobic substrates was in general higher than BSA in terms of
numbers of molecules attached (Fig. 2, bottom panel), reflecting the
smaller size and consequently reduced "footprint" of this protein
on the surface.
Adhesion of microorganisms. Having established satisfactory protocols for surface modification and proven that the surfaces displayed sufficiently high density of the graft material even on a molecular scale, we proceeded to investigate the adhesion of microorganisms. L. monocytogenes, S. typhimurium, S. aureus, and E. coli were chosen as representative pathogens, although most of the initial experiments were conducted with L. monocytogenes. Interestingly enough, it was found that the pattern of L. monocytogenes adhesion was not too dissimilar from that of BSA. Once again, the greatest number of cells attached to hydrophobic substrates and the charged APTES surface. The hydrophilic surfaces of MeO-PEO-3 and MeO-PEO-5000 showed the highest resistance to cell attachment, according well with previous studies of these materials (25). As before, the acetamide graft surfaces also displayed low cell adhesion, although the long-chain analog poly(N-methylacrylamide) was less effective (Fig. 3). MeO-PEO-5000-modified substrates were found to be the least adsorptive for other bacteria, too, but the hydrophilic acetamide surface was less effective at preventing the adhesion of S. aureus and S. typhimurium than the low-surface-energy hydrophobic Krytox perfluoropolymer (Fig. 4). This result indicates that, for materials which are not particularly repellent for microorganisms, the surface chemistry of bacteria themselves plays an important role in the adsorption process. It is also worth noting that the kinetics of microorganism attachment were very different from those observed with proteins; in general, the number of cells bound to the substrates reached a stable level after 16 to 24 h of incubation compared to 1 h or less for proteins.
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1), suggested that significant amounts of exudates
might have been present.
We therefore attempted to check the possibility of prior exudate
adsorption by looking at the competition between bacterial exudates and
radioactively labelled BSA for representative substrates. To this end,
the derivatized coverslips were incubated with supernatant obtained
from bacterial cultures, and the adsorption of BSA to these pretreated
substrates was tested as described above. It was found that for
hydrophobic (perfluoroalkyl) and charged (aminopropyl) surfaces, BSA
adsorption was partially suppressed by 4 to 23%, but for the
hydrophilic surfaces (acetamide) it actually increased by 40%,
although the actual amount of BSA adsorbed in the latter case was
still low (0.7 pmol · cm2). Thus, it seems that
these differences are not sufficient in themselves to
support the hypothesis that L. monocytogenes excretes specific macromolecules in solution to facilitate cell adhesion. However, this may well be the case for other microorganisms, such as
staphylococci, which are known to attach to hydrophobic surfaces much
more readily. Further experiments are currently being conducted to
elucidate this possibility.
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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The prerequisite for our study of bacterial adhesion was the preparation and characterization of suitable, well-defined substrates such that the effects of physicochemical parameters, including hydrophobicity, hydrophilicity, steric hindrance, roughness, graft density, and functionality, on the adsorption phenomena could be assessed individually. The synthetic substrates (Table 1) were shown by microscopy and contact angle goniometry to be smooth over a "sub-bacterial" scale (i.e., <1 µm in length). Some evidence for surface rearrangements on APTES substrates was obtained during dynamic and equilibrium contact angle measurements, as judged by the hysteresis observed between the advancing and receding contact angles. The most probable explanation for this was that as the water drop advanced over the surface, amine groups were able to rearrange their conformation to point into the aqueous layer. Subsequent removal of the water drop left a more hydrophilic surface, which reverted to a more hydrophobic structure as amine groups rebound to the silica surface. However, all of the other substrates appeared to be stable and homogeneous at 37°C. The experimentally determined surface energies for derivatized substrates were as expected for materials with a high graft density, and this, combined with the microscopy (AFM and optical) and functional-group titration data, indicated that the substrates were chemically well defined and thus suitable as probes for studying adsorption phenomena.
Further evidence for the satisfactory surface density of the introduced
functionality on a macromolecular scale came from the protein
adsorption studies. BSA adsorption to the aminopropyl surfaces was
close to the calculated figure for a monolayer (we assumed a surface
area for BSA of 1.0 × 1016 m2, based
on approximate dimensions of 100 by 100 Å, and considered that no
denaturation took place) and was essentially irreversible; increasing
the amount of BSA in the medium did not lead to further adsorption. For
cytochrome c the highest adsorption occurred to a
hydrophobic uncharged surface and represented a coverage of ~90% of
a monolayer, again assuming no denaturation of the protein took place
on attachment. For "noncharged" alkylamide grafts, a clear
correlation was obtained between the chain length of the amide and the
numbers of protein molecules attached, with BSA adsorption reaching a
maximum at alkyl chains containing six to eight methylene units. Thus,
it appeared that with these and higher homologs, adsorption to what was
effectively a hydrocarbon layer occurred. Attachment of the proteins to
other hydrophobic substrates was as expected, with low-surface-energy
perfluorinated materials displaying adsorption similar to that of the
long-chain hydrocarbons.
The effect of grafting hydrophilic groups to the substrates was largely
in accordance with previous studies (15). The hydrophilic MeO-PEO-5000 surface displayed the best protein-rejecting properties, and the short-chain MeO-PEO-3 also exhibited low protein adsorption (20, 18), although this latter surface proved less resistant to the smaller cytochrome c. The amine-terminal APTES
surface might also have been expected to be hydrophilic under the
experimental conditions (pH 7.0, 37°C); however, the surface energy
recorded (34.76 mJ · m2) was similar to that of
the longer-chain alkylamides. This suggested that at least some of the
amine groups may have been "folded back" by binding to the siloxane
surface, exposing the propyl chains to the solution (36).
Reaction of the APTES substrates with acetyl chloride generated a more
hydrophilic acetamide surface (39.38 mJ · m2), and
this in turn reduced protein adsorption to levels similar to those of
the MeO-PEO-3 substrates. The polymer analog of acetamide, poly(N-methylacrylamide) (PNMeAM) exhibited a rather higher
degree of adsorption of BSA than did cytochrome c, which was
not expected since it is hydrophilic, of high surface energy (40.66 mJ · m2), and might, in accordance with the
mechanisms advanced for protein rejection by PEO, offer steric
hindrance to attaching moieties via polymer chains extending into
solution (indeed, MeO-PEO-5000 proved more resistant to cytochrome
c adsorption than its short-chain counterpart MeO-PEO-3 in
our experiments). It seems, therefore, that a simple combination of
hydrophilicity and steric hindrance is not necessarily enough to reduce
protein attachment, and the fact that MeO-PEO-5000 adsorbed far less
BSA than PNMeAm perhaps indicates that it is the unique solution
structure of PEO polymers (39), rather than their
hydrophilicity and molecular weight, which accounts for their protein-
and microorganism-rejecting abilities.
Bacterial adhesion was also lowest to the hydrophilic substrates
MeO-PEO-3 and MeO-PEO-5000, although the numbers of cells attached to
these substrates varied markedly between different microorganisms
(e.g., 8.6 × 104 cells · cm2 for
S. typhimurium on MeO-PEO-5000 and 2.3 × 106 cells · cm2 for S. aureus). In addition, there was a considerable difference in the
attachment of different bacteria to the hydrophilic acetamide surface.
Whereas the adhesion of L. monocytogenes and E. coli to acetamide followed a pattern similar to that of BSA and
cytochrome c, attachment of S. typhimurium and
S. aureus was higher to this substrate than to the
hydrophobic Krytox surface. This may have been due to the latter
bacteria exuding adhesion-promoting materials which adsorbed
sufficiently to mask the underlying hydrophilic surface. In this case,
adsorption of much less than a monolayer of exudate might result in
sticky "patches" on the surface that are separated but within
"bridging" distance for a cell. As a corollary, for the same
hydrophilic acetamide surface and L. monocytogenes, preadsorption of BSA suppressed cell attachment considerably, even
though the amount of BSA was ~10% of a monolayer.
Admittedly, such a consideration treats the microorganisms as "living colloids" (5), disregarding the specific roles of bacterial structures such as pili, cell wall components, and extracellular lipopolysaccharides which have been the subject of much work elsewhere by a number of groups (21, 28) and which are considered to be of particular importance in the later stages of biofilm formation. However, it has been argued by a number of authors that these structural features are of less significance in the initial stages of the attachment process than the intrinsic thermodynamic factors involved (4, 29), and a number of detailed studies have been carried out to support this assertion (37). Our study has shown that over the time period of protein adsorption and initial cell attachment (1 to 24 h), this assumption is reasonable, since the overall pattern of cell adhesion to different substrates was similar among the various cell types, although the absolute numbers varied considerably.
In conclusion, hydrophilic uncharged surfaces showed the greatest resistance to protein adsorption and cell attachment, and a clear correlation between substrate chemistry and protein adsorption was established. In addition, our studies supported the view that the effectiveness of PEO polymers in preventing protein adsorption cannot be attributed directly to hydrophilicity and molecular weight alone. For our model systems, protein adsorption could be approximately correlated with short-term cell adhesion, and bacterial attachment for some microorganisms also correlated with substrate chemistry. However, for S. aureus and S. typhimurium a different mechanism of cell attachment appeared to occur, as shown by the relative numbers of cells adsorbed to hydrophobic and hydrophilic substrates. Preadsorption of BSA resulted in a reduction of cell adhesion to selected surfaces in 24-h assays. The implications of this for in vivo attachment of cells suggest that hydrophilic passivating groups, if grafted uniformly and in sufficient density to the surface, may be the best method for preventing cell adsorption to synthetic substrates.
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ACKNOWLEDGMENTS |
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This work was funded by the Ministry of Agriculture Fisheries and Food.
We would like to thank Frans Llevat and Laura Magraner for help with some of the experimental work; John Tsibouklis, Adrian Thorpe, and Simon Young, University of Portsmouth, for contact angle measurements; and Terry Roberts (Ministry of Agriculture, Fisheries, and Food) for many helpful discussions.
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FOOTNOTES |
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* Corresponding author. Mailing address: Institute of Food Research, Norwich Research Park, Colney, Norwich, NR4 7UA, United Kingdom. Phone: (44) (0) 1603 255000. Fax: (44) (0) 1603 507723. E-mail for C. Alexander: cameron.alexander{at}bbsrc.ac.uk. E-mail for E. N. Vulfson: jenya.vulfson{at}bbsrc.ac.uk.
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REFERENCES |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Applied and Environmental Microbiology, November 1999, p. 4995-5002, Vol. 65, No. 11
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