ARTICLE pubs.acs.org/Langmuir
Probing Roles of Lipopolysaccharide, Type 1 Fimbria, and Colanic Acid in the Attachment of Escherichia coli Strains on Inert Surfaces Yuanqing Chao and Tong Zhang* Environmental Biotechnology Laboratory, The University of Hong Kong, Pokfulam Road, Hong Kong SAR, China
bS Supporting Information ABSTRACT: The roles of bacterial surface polymers in reversible (phase I) and irreversible (phase II) attachment (i.e., lipopolysaccharides (LPS), type 1 fimbria, and capsular colanic acid (CA)) were investigated in situ by combining fluorescence microscopy and atomic force microscopy. Fluorescence microscopy was used to evaluate the phase I attachment by counting the total number of cells on the substrata, and AFM was applied to image the phase II cells and measure the lateral detachment force to characterize phase II attachment. Also, by comparing the number of cells in phases I and II, the transformation ratio was calculated and used as an index to evaluate the roles of different polymers in the attachment process. Escherichia coli K-12 and its six mutants, which had different surface polymers in terms of LPS structures, CA contents, and type 1 fimbriae, were used as the test strains. Six different materials were applied as substrata, including glass, two metals (aluminum and stainless steel), and three plastics (polyvinyl chloride, polycarbonate, and polyethylene). The results indicated that LPS significantly enhanced phases I and II attachment as well as the transformation ratio from phase I to II. Like LPS, type 1 fimbriae largely increased the phase I attachment and the transformation ratio; however, they did not significantly influence the adhesion strength in phase II. CA had a negative effect on attachment in phases I and II by decreasing the adhered number of cells and the lateral detachment force, respectively, but had no influence on the transformation ratio.
1. INTRODUCTION In many natural and artificial environments, bacteria may attach to surfaces and subsequently form biofilms.1 It is widely accepted that the biofilm formation process has several major phases,2 including (I) initial reversible attachment, (II) a transition to irreversible attachment, (III) biofilm architecture development, (IV) mature biofilm formation, and (V) cell dispersion. Because phase I and II attachments are the crucial and limiting steps in the development of the biofilm, many studies have focused on these two phases in the attachment of bacterial cells to various surfaces.3 There are several distinguishing physical and chemical properties between the two phases of single-cell attachment.4 In phase I attachment, bacterial cells are transported or swim by themselves close enough to the substrata and then initial reversible attachment occurs because of van der Waals forces, electrostatic forces, and hydrophobic interactions.1,5,6 Subsequently, attached cells stick themselves on the surface by producing extracellular polymeric substances (EPS) that allow them to transfer to phase II attachment.7 In the transformation from phase I to II attachment, other stronger interactions (e.g., covalent and hydrogen bonds) and hydrophobicity interactions are involved.4 Previous studies proved that various cell-surface polymers, such as lipoplysaccharide (LPS) and filamentous proteins (fimbriae and flagella) in the outer membrane as well as capsular colanic acid (CA) surrounding the outer membrane for many r 2011 American Chemical Society
bacterial strains, could influence the bacterial attachment process on hosts or inert surfaces.2 Various methods have been applied to evaluate the roles of surface polymers during the bacterial attachment process. Several studies evaluated the role of LPS and extracellular polymeric substances in bacterial adhesion by using a filtration system in packed columns.8,9 However, the experimental columns were not well-defined in terms of hydrodynamic flow conditions and allowed only the macroscopic observation of bacterial attachment.10 To observe and evaluate the roles of surface polymers during the attachment process on the microscopic level, many studies applied various microscopes in their examination, including a light microscope,11 a fluorescence microscope,12,13 a confocal laser scanning microscope (CLSM),14 a scanning electron microscope (SEM),1517 and a transmission electron microscope (TEM).12,15 These studies provided quantitative information about the adhesion number of cells, biofilm coverage, and 3D biofilm structure and gave many critical insights into the roles of surface polymers during the bacterial attachment process and biofilm development. Unfortunately, the above quantification methods are not able to separate phases I and II in single cell attachment and provide information about the adhesion force that is crucial in phase II. Received: March 18, 2011 Published: August 15, 2011 11545
dx.doi.org/10.1021/la202534p | Langmuir 2011, 27, 11545–11553
Langmuir
ARTICLE
Figure 1. Scanning electron microscopy images of E. coli strains (A) K-12 and (B) JW4277, which were taken at magnifications of 20 000 and 10 000, respectively. Arrows in image A indicate the type 1 fimbriae. (C) Capsular colanic acid production of E. coli strains JW2039 and JW5917. The CA contents were converted to glucose by the phenolsulfuric acid test. Data are the means (three replicates) ( the standard error.
The cellsubstrata adhesion force has been largely investigated by using shear forces to detach the adherent bacterial cells and measure the lateral detachment force to characterize phase II attachment (e.g., flow chambers,18 spinning disks,19 centrifugation,20 and air bubbles21). However, these methods are limited by several major disadvantages, including low sensitivity and precision, complicated and time-consuming sample-preparation procedures, the inability to apply both controlled and directed forces at a specific location, and difficulty in estimating the critical force needed to detach all of the adhered cells.3,17,22 Recently, atomic force microscopy (AFM) has also been used to determine the lateral detachment force between adherent cells and substrata in situ because AFM can scan samples on the nanometer or subnanometer scale and probe the interactions of cells and the substratum in adhesion processes with high force resolution in either dry or wet environments.23 Senechal et al. evaluated the force required to detach Enterococcus faecalis cells from polymer substrata according to the calibration curve of the deflection set point applied force via AFM and demonstrated an accurate approach to determining the adhesion forces for bacterial phase II attachment.17 Other studies evaluated the lateral detachment force of bacteria according to the total compression of the cantilever, probe geometry, and cantilever orientation by using the contact mode of AFM and more accurately measured the lateral detachment forces of cells.3,22 These studies demonstrated that the contact mode of AFM could be used to quantify the lateral force required to detach adhered bacterial cells from the substratum accurately and rapidly. However, studies in which AFM is applied to evaluate the roles of bacterial surface polymers in phase II attachment are rare. To investigate the roles of LPS, type 1 fimbriae, and CA in phases I and II of single-cell attachment comprehensively, in the present study the fluorescence microscope was used to evaluate the phase I attachment by counting the total number of cells on the substrata, and AFM was applied to image the phase II cells
and measure the lateral detachment force to characterize the phase II attachment. Also, by comparing the number of cells in phases I and II, the transformation ratio was calculated and used as an index to evaluate the roles of different polymers in the attachment process.
2. MATERIALS AND METHODS 2.1. Bacterial Strains and Growth Conditions. E. coli strain K-12 BW25113 and six E. coli K-12 mutants, including D21f2, D21, JM109, JW2039, JW4277, and JW5917, were obtained from the E. coli Genetic Stock Center (Department of Biology, Yale University) and used in this study as the test strains. D21f2, D21, and JM109 were widely used in previous studies on bacteria attachment10,13,24,25 because they have different lipopolysaccharide (LPS) outer layers. E. coli JM109 had an entire LPS, including a ketodeoxyoctonate (KDO), an inner and outer core polysaccharide, and a relatively larger O-antigen. The LPS of D21 consisted of the former two parts but no O-antigen. The LPS of D21f2 contained only KDO in the outer membrane. These three strains were applied to evaluate the roles of bacterial LPS structure during single-cell attachment. Other mutants, including JW2039, JW4277, and JW5917, are isogenic strains, and their parent is BW25113 strain. E. coli JW4277 does not have the fimA gene, which is responsible for the production of proteins called fimbrins. These fimbrins made up the core of the fimbrial rod and the fibrillar tip of the type 1 fimbriae.26 The data in our pre-experiment (Figure 1A,B) showed that the type 1 fimbriae production of JW4277 largely decreased compared with its parent strain, K-12 BW25113. These two isogenic strains were used to study the roles of type 1 fimbriae in single-cell attachment. The two mutants, JW2039 and JW5917, have different capsular colanic acid (CA) contents (Figure 1C). The CAs of JW2039 and JW5917 were extracted, purified, and quantified according to a previous study.27 For JW2039, the CA synthesis gene wcaF was knocked out, and its CA production was significantly inhibited, whereas for JW5917, the regulator of CA production gene rcsC was deleted and the CA production is more than 11546
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir
ARTICLE
that of JW2039. These two isogenic mutants were applied to determine the roles of CA during single-cell attachment. The strains were cultivated at 37 °C and 150 rpm in a sterilized (121 °C for 20 min) LuriaBertani (LB) medium that contained 10 g of tryptone, 5.0 g of yeast extract, 5.9 g of NaCl (controlling the ionic strength of the medium to be 100 mM), and 1 L DI water, and the pH was adjusted to 7.4. Then the cells were harvested in the log phase at a concentration equivalent to an optical density at a 600 nm (OD600 nm) value of about 0.2 (approximate 107 CFU/mL; CFU stands for colony-forming unit). These bacterial solutions were then immediately used for further experiments. 2.2. Substrata Preparation. Six different inert materials were used as substrata in this study, including glass, two metals (stainless steel and aluminum), and three plastics (polyvinyl chloride (PVC), polycarbonate (PC), and polyethylene (PE)). All of the substrata were cylindrical with a diameter of 10 mm and a height of 2 mm. The stainless steel and aluminum were polished with a Micropolish Alumina solution with a particle size of 0.3 μm (no. 40-6352, Buehler, USA). Afterward, all of the substrata were washed by sonication for 10 min in DI water and then in dehydrated ethanol (Merck KGaA, Germany). Then the substrata were dried under a flow of nitrogen, glued onto glass-slides using Crystalbond 209 (Ted Pella, USA), and stored in a desiccator at room temperature (23 ( 1 °C) before use. 2.3. Bacterial Adhesion. An O-ring was glued onto the glass slide by using high-vacuum grease (Dow Corning Corporation, USA) to hold the solution to provide an in situ environment during measurements. First, 2 mL of a prepared bacterial solution was added to the O-ring, and the surface of the prepared substratum was immersed and cultivated at room temperature for 1 h. The samples were carefully washed three times with sterilized 100 mM PBS solution (0.29 g of KH2PO4, 1.19 g of K2HPO4, and 4.93 g of NaCl in 1 L of DI water, pH adjusted to 7.2, and sterilized at 121 °C for 20 min). Then the samples were immersed in the above PBS solution and placed in a humid chamber at room temperature for 12 h before AFM measurement. A previous study showed that the lateral detachment force of adhered cell adhesion would be stable during the following AFM measurements (35 h) after 13 h of cultivation (1 h in LB + 12 h in PBS).3 2.4. AFM Measurement. AFM images were obtained in contact mode using JPK NanoWizard AFM (JPK Instruments, Germany) and used to calculate the lateral detachment force of adherent E. coli bacterial cells on substrata. The quantitative method for determining the lateral detachment force was described in our previous study.3 Briefly, considering the total compression of the cantilever, probe geometry, cantilever orientation, and bend during the bacterial cell detachment event, the lateral detachment force could be calculated from eq 1: Flat ¼ kSVtotal sinðΦ þ θÞ 8 2 qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi39 > > < L ðVtotal SÞ2 þ ðL cos ΦÞ2 7= 6 cos Φ þ θ 2 arctan4 5 > > Vtotal S þ L sin Φ : ;
ð1Þ In eq 1, the cantilever’s spring constant k and sensitivity S were calibrated by the resonance frequency of the cantilever and the slope of the force spectrum between the tip and substratum using software provided by the manufacturer, respectively. Angles Φ and θ were parameters of the cantilever orientation and probe geometry. L presented the length of the applied cantilever. For the AFM system used in this study, θ = Φ = 10° and L = 300350 μm. Vtotal was the total compression of the cantilever and could be determined by the sum of Vset point and Vdeflection. Vset point was the user-defined cantilever compression for each scan. Vdeflection showed an additional compression of the cantilever during the bacterial detachment event.3,22 A scan size of 40 40 μm2 and scan rate of 0.5 Hz were used in the lateral detachment force measurements.3 Silicon cantilever CSC38
(Mikromasch, Estonia) with a series of spring constants of 0.03, 0.05, and 0.08 nN/nm was applied for force measurements. Before the AFM determination, the spring constant and sensitivity of each cantilever were recalibrated on a clean glass slide. At the start of each measurement, the sample was scanned under the minimum force (0.21 ( 0.04 nN) over a random area. Then the applied force was gradually increased until all of the observed adherent cells were detached from the surface. For each bacterium/substratum combination, 12 individual cultures were conducted, 58 fields were randomly selected, and 10100 cells were measured. All measurements were conducted in the PBS solution. Besides the lateral detachment force measurements, AFM was also used in tapping mode to characterize the morphology of adhered E. coli cells on several substrata in situ. Silicon cantilever CSC37 (Mikromasch, Estonia) with a series of spring constants of 0.3, 0.35, and 0.65 nN/nm was applied for morphology characterization in 100 mM PBS solution. To decrease the applied force between the cantilever tip and bacteria to minimize the influence on the bacterial morphology during AFM scanning, the set-point value of the oscillation amplitude was maintained at a higher value (1.2 V or above) than the free amplitude of the cantilever (normally 1.0 V). Measurements were conducted by scanning several random areas with the scan size of 20 20 μm2 at a scan rate of 0.2 Hz. 2.5. Total Cell Counting and Viability Test. The live/dead stain method was used here to count the total number of cells in phase I and II attachment and to evaluate the bacterial viability.28 A LIVE/ DEAD Bac Light bacterial viability kit (lot 52027A, Molecular Probes Inc., Eugene, OR) was used in the present study. The kit contains fluorescent nucleic acid stains SYTO9 for live cells and propidium iodide for dead cells. After AFM measurement, 5 μL of the kit was added to the substrata surface. After 10 min of staining, eight fields were randomly selected and photographed with a fluorescence microscope (Nikon Eclipse E600, Japan) equipped with a 100 objective. Then the total number of cells (number of live cells + number of dead cells) and viability ((no. of live cells/total number of cells) 100%) were counted and calculated according to these images, respectively. 2.6. Hydrophobicity of Bacteria and Substrata. To evaluate the bacterial and substrata hydrophobicity, contact angle measurements were performed using a contact angle measurement machine (Powereach, Shanghai) by applying the sessile drop method.29 One drop of a liquid (35 μL) was dripped onto a prepared substrata surface or a lawn of bacteria. The images were photographed in 1 s, and the contact angle was measured using Microsoft Office Visio (2003). Lawns of bacteria were prepared by filtering 20 mL of a cell suspension (108 CFU/mL, prepared as described above) onto a 0.2 μm MCE filter (GSWP09000, Millipore Corporation). After filtration, the filter was dried in a Petri dish for at least 30 min at room temperature and measured within 1 h. Previous studies have shown that after a drying period of 30 min, measurements on bacterial lawns are stable for several hours.13,29 Contact angles were measured by using one nonpolar (diiodomethane) solution and two polar (formamide and DI water) solutions. All experiments were conducted for six replicates (two samples three measurements). The hydrophobicity of tested strains and substrata was evaluated by the surface tension (γ) equilibrium,30 which is expressed by eq 2 (Young’s equation): pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi γ ¼ γLW þ 2 γABþ γAB ð2Þ Superscripts LW and AB represent the surface tensions originating from Lifshitzvan der Waals and Lewis acidbase interactions, respectively. Symbols “+” and “” stand for the electron acceptor and donor, respectively. The relationship between the contact angle (θ) and the surface tension components was described by eq 3 (YoungDupre equation) qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi LW þ γABþ ð1 þ cos θÞγL ¼ 2ð γLW γAB γABþ þ γAB Þ L L S S γL S ð3Þ 11547
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir
ARTICLE
Table 1. Surface Tension (mJ/m2), Zeta Potential (mV) of E. coli Strains and Substrata (Ionic Strength = 100 mM), and rms Roughness (nm) of Six Substrata surfaces strains
substrata
D21f2
surface tension
zeta potential
53 ( 0.89
31 ( 4.6
roughness
Table 2. Bacterial Viabilities (%) of Seven E. coli Strains on Six Substrata D21f2
D21
JM109 K-12 JW4277 JW2039 JW5917
glass
95 ( 2 91 ( 1 86 ( 0 92 ( 5 98 ( 1
91 ( 3
93 ( 2
PVC PC
75 ( 4 97 ( 0 95 ( 3 89 ( 2 93 ( 2 93 ( 3 97 ( 2 92 ( 2 91 ( 1 88 ( 3
97 ( 1 84 ( 1
85 ( 3 97 ( 0
D21
45 ( 1.5
25 ( 8.3
JM109
38 ( 2.2
19 ( 5.3
PE
95 ( 0 92 ( 1 89 ( 2 87 ( 3 95 ( 1
98 ( 0
91 ( 3
K-12
44 ( 1.3
21 ( 4.4
aluminum
90 ( 2 95 ( 1 91 ( 2 91 ( 4 94 ( 0
96 ( 3
98 ( 0
JW4277 JW2039
50 ( 1.1 44 ( 2.1
25 ( 2.8 16 ( 5.7
stainless steel 98 ( 1 96 ( 1 83 ( 3 94 ( 1 89 ( 4
97 ( 0
98 ( 0
JW5917
56 ( 0.76
34 ( 6.7
glass
57 ( 0.70
26 ( 4.9
0.64 ( 0.091
PVC
36 ( 1.1
38 ( 2.1
5.5 ( 1.9
PC
42 ( 1.1
27 ( 4.6
21 ( 8.5
PE
34 ( 1.6
29 ( 2.3
21 ( 6.4
aluminum
44 ( 0.65
20 ( 6.4
4.4 ( 1.2
stainless steel
41 ( 1.8
40 ( 7.3
2.1 ( 0.54
where subscripts S and L represent the surface and the liquids that were used, respectively. 2.7. Surface Charge of Bacteria and Substrata. The zeta potentials of bacteria, assumed to be equal to the cell-surface charge at high ionic strength,31 were calculated from electrophoresis mobility by using the Smoluchowski equation.32,33 Cells prepared as described above were resuspended in 100 mM PBS (relatively high ionic strength), and zeta potentials were measured 5 times using 10 cycles per analysis (Delsa Nano Zeta Potential Analyzer, Beckman Coulter). The zeta potentials of the substrata surface were determined after fixing the coupon in a flow cell and measuring with the same equipment. All determinations were conducted for three replicates. 2.8. Roughness of the Substrata. The roughness of six substrata surfaces was determined by using the contact mode of AFM under an air environment. A scan size of 2 2 μm2 was used to measure the surface roughness because the roughness on the bacterial scale could be more representative for evaluating its influence. Height images were applied to calculate the roughness based on root-mean-square (rms) values. Ten fields were randomly selected and measured for each substratum.
3. RESULTS 3.1. Physiochemical Properties of Bacterial and Substrata Surfaces. The physiochemical properties of tested E. coli strain
surfaces, including the hydrophobicity and surface charge (Table 1), were characterized by the contact angle (Table S1) and zeta potential, respectively. For strains with different LPS structures, the cells (JM109) with full LPS were more hydrophilic (PJM109-D21 < 0.001, PJM109-D21f2 < 0.001) and less negatively charged (PJM109-D21 = 0.159, PJM109-D21f2 = 0.005) than others (D21 and D21f2) with partial LPS structures. Comparing E. coli K-12 with type 1 fimbriae-lacking strain JW4277, there were significant but slight differences in their surface tensions (P < 0.001) and charges (P = 0.129), indicating that the presence of type 1 fimbriae could make cells slightly more hydrophobic and positively charged. For two other strains JW2039 and JW5917, the CA made cells more hydrophilic (P < 0.001) and more negatively charged (P = 0.001). For the applied substrata, the glass was the most hydrophilic, followed by metals and plastics according to the contact angles and total surface tensions (Table 1 and S1). The zeta potential values showed that the plastics were more negatively charged
Figure 2. (A) Fluorescent image and (B) AFM vertical deflection image of adherent E. coli K-12 cells on polished stainless steel (scale bar = 4 μm). (A) The fluorescent image shows the total number of cells obtained by the live/dead staining method. (B) The AFM image presented the attached cells in phase II only.
(Pglass-PC < 0.001, Pglass-PE = 0.237) than glass except for PVC (Table 1). A huge difference appeared between the zeta potential values of two metals. This might be mainly due to the quick oxidation of the aluminum surface and the fact that the stainless steel was much more stable during the experiment. The surface roughness measurements suggested that glass had the smoothest surface, followed by polished metals and plastics (Table 1). 3.2. Bacterial Viability. The live/dead staining results (Table 2) showed that all E. coli strains maintained a high viability (averaging 92%, ranging from 75 to 98%) after 20 h (13 h of cultivation + 7 h of AFM operation in an LB medium and a PBS solution) at room temperature. This indicated that the test strains did not lose their viability during measurements and the obtained results can reflect the real situations during bacterial attachment events. 3.3. Determination of the Transformation Ratio from Phase I to II Attachment. A phenomenon was observed during the experiment in which the total number of adherent cells counted under the fluorescence microscope was much higher than that obtained by AFM imaging (Figure 2). The total number of cells reflected the amount of all of the adherent cells in either phase I or II attachment (Figure 2A). However, the number of cells obtained by AFM were only for those in phase II attachment because the phase I cells were removed easily during AFM scanning and were not able to be detected by cantilever and displayed in AFM images (Figure 2B). Moreover, because all of the phase II cells were transferred from phase I, the total number of cells could be used as an indicator of phase I attachment. Therefore, the ratio of the AFM number of cells to the total number of cells can be used as an index to reflect the transformation percentage from phase I to II attachment (Table 3). 11548
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir
ARTICLE
Table 3. Transformation Ratio (%) from Phase I to II Attachment of Seven E. coli Strains on Six Substrata D21f2
D21
JM109
K-12
JW4277 JW2039 JW5917
glass
34 ( 5 36 ( 6 44 ( 5 39 ( 7
28 ( 3
45 ( 7
51 ( 3
PVC
53 ( 7 49 ( 5 54 ( 2 56 ( 2
14 ( 2
33 ( 3
43 ( 4
PC
13 ( 3 19 ( 5 50 ( 8 54 ( 6
19 ( 1
32 ( 2
24 ( 6
PE
23 ( 4 19 ( 5 31 ( 4 20 ( 5
17 ( 3
18 ( 2
25 ( 5
aluminum
40 ( 4 48 ( 2 49 ( 3 55 ( 4
38 ( 4
32 ( 3
27 ( 7
stainless steel 26 ( 7 35 ( 2 38 ( 3 50 ( 3
26 ( 6
22 ( 3
26 ( 5
Figure 4. (A) Total number of cells and (B) lateral detachment force of type 1 fimbria mutant JW4277 and E. coli K-12 BW25113 on six substrata. The error bars represent standard errors of the means. An * beside a letter indicates that the mean value is significantly different (at the 5% level).
Figure 3. (A) Total number of cells and (B) lateral detachment force of LPS mutants D21f2, D21, and JM109 on six substrata. The error bars represent the standard errors of the means. An * beside a letter indicates that the mean value is significantly different (at the 5% level).
3.4. Role of LPS in the Attachment of E. coli. Three E. coli mutants (i.e., D21f2, D21, and JM109) were used in the present study to evaluate the role of LPS in the attachment process. The total number of cells (Figure 3A) counted by live/dead staining showed that there were no significant differences between D21f2 and D21 on six substrata, indicating that the inner and outer core polysaccharides played less-significant roles in the phase I attachment. The transformation ratios shown in Table 3 implied that the inner and outer core polysaccharides of LPS did not significantly influence the transformation ratio from phase I to II either. And the lateral detachment forces of these three strains measured by AFM were also similar as shown in Figure 3B, indicating that the inner and outer core polysaccharides did not enhance the adhesion strength between the bacteria and substrata in phase II. However, the O-antigen of LPS might play an important role during the single-cell attachment process of E. coli. Mutant JM109 containing a relative larger O-antigen, compared
with D21, had significantly more adherent cells than those of D21f2 and D21 in phase I (Figure 3A), higher transformation ratios from phase I to II (Table 3), and a higher lateral detachment force in phase II (Figure 3B). 3.5. Role of Type 1 Fimbria in the Attachment of E. coli. Strain K-12 and mutant JW4277 of E. coli were compared to study the role of type 1 fimbria in the bacterial attachment process. The results showed that the type 1 fimbriae significantly increased the total number of cells in phase I (Figure 4A) and the transformation ratios (Table 3). However, the lateral detachment forces of cells in phase II on substrata (Figure 4B) were not significantly influenced by the type 1 fimbriae except on stainless steel. 3.6. Role of CA in the Attachment of E. coli. Two mutants JW2039 and JW5917, which contained significantly different contents of CA, were used to study the role of CA in the attachment process. The results showed that the CA content inhibited the phase I attachment significantly except on glass (Figure 5A) but had no impact on the transformation ratios (Table 3). Moreover, judging from the lateral detachment force (Figure 5B), CA significantly decreased the adhesion strength of E. coli mutants on five of six substrata in phase II.
4. DISCUSSION 4.1. Influence of Interfacial Physicochemical Properties on the Phase I Attachment. The phase I attachment (reversible
attachment) was considered to be based on the physicochemical interactions at the interfaces of bacteria and substrata.34 Previous 11549
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir
Figure 5. (A) Total number of cells and (B) lateral detachment force of E. coli mutants JW2039 and JW5917 on six substrata. The error bars represent the standard errors of the means. An * beside a letter indicates that the mean value is significantly different (at the 5% level).
studies were conducted to evaluate the bacterial attachment process in terms of the hydrophobicity and surface charge of bacteria and substrata,13,35 solution chemistry including the ionic strength and pH,9,35 and the roughness and microtopography of the substrata.3638 However, it is difficult to investigate the attachment process on the basis of single or double interfacial parameters because bacterial attachment to the substratum is a multifactor process and other surface properties may also play significant roles in the attachment.1 To understand the roles of physicochemical properties during the bacterial attachment process comprehensively, cluster analysis and canonical correspondence analysis (CCA) were conducted in the present study. (For details, please refer to the Supporting Information.) Cluster analysis showed that the grouping pattern of the phase I attachment was related to the involved physicochemical properties. The results (Figure S1) illustrated that the phase I attachment was significantly grouped on the basis of the substrata, especially for glass and stainless steel. However, the phase I attachment on plastics and aluminum were not clearly grouped. This pattern might be caused by the similar physicochemical properties of these substrata (Table 1). The results of CCA (Figure S2) showed that the phase I attachment had positive correlations with the hydrophobicity and surface charge of bacteria and with the surface charge of the substrata but no significant relationship with the hydrophobicity and rms roughness of substrata. This indicated that the physicochemical properties caused by different bacterial surface properties (e.g., LPS, fimbriae, and colanic acid) could significantly influence the bacterial phase I attachment. Moreover, CCA also
ARTICLE
revealed that the substrata surface charge may play significant roles during phase I attachment. This was consistent with the observation that the phase I number of cells on stainless steel was significant more than that on other substrata (Figures 3A, 4A, and 5A). Because the stainless steel had a positive surface charge of 40 mV in 100 mM PBS whereas other substrata had a negative charge that varied from 20 to 38 mV (Table 1), this indicated that the electrostatic force between bacteria and stainless steel may be much larger than other combinations and may become the dominant factor in the phase I attachment. 4.2. Characterization of Adhesion Strength in Phase II by the Lateral Detachment Force. To characterize the adhesion strength of phase II attachment, the lateral detachment force tests were conducted by AFM in the present study. A previous study indicated that after bacterial cells stuck themselves on the surface by the surrounding EPS in phase II, the strength and number of chemical bonds between bacteria and the surface, rather than the surface physicochemical properties, would be the dominant factor during this phase.7 The bacterial morphology characterization in the present study showed that the phase II cells were firmly adhered to the substrata by the surrounding EPS (Figure 6). The height information in the cross section also illustrated that the amount of EPS was relatively large because the height of EPS was identical to that of bacterial cells (Figure 6B). The results of lateral detachment force measurements showed that E. coli strains adhered the most firmly to the plastic surface, followed by metals and glass (Figures 3B, 4B, and 5B). This revealed that the short-range chemical interactions (ionic, hydrogen, and covalent bonds) between bacteria and plastics were more complex or stronger than those between bacteria and metals. Another possible explanation is related to the roughness of the applied substrata. The measured lateral detachment force was significantly correlated with the rms roughness of the substrata (Figure 7). For a rougher substratum, the contact area between the bacterial cell/EPS and the substratum surface was larger than that on a smooth substratum. This can significantly increase the number of chemical bonds, and the cell may be glued more firmly to the surface. This suggests that the roughness might play an important role in the phase II attachment. The CCA analysis (Figure S2) also indicated that the phase II attachment had a positive correlation with the rms roughness of the substrata. Several studies have been determined the lateral detachment forces of various cell/substratum combinations using the contact mode of AFM. Senechal et al. measured the lateral detachment forces of E. faecalis on three polymers.17 The results indicated that the detachment forces varied from 0.7 to 19 nN depending on the substratum. Deupree and Schoenfisch found that the detachment forces needed to remove Staphylococcus aureus from a xerogel film increased from 4 to 8 nN over 10 h of cultivation.22 Moreover, He et al. obtained an even larger lateral detachment force (>80 nN) of adherent S. aureus cells on glass coated with adhesive ligands.39 In the present study, the measured lateral detachment forces of adherent E. coli cells on six substrata were relative smaller and varied from 0.3 to 1.2 nN. These findings indicated that the bacterial detachment forces were strongly dependent on the bacterial species, substratum, cultivation procedures, and time. The large difference between the previous studies and the present study might be mainly due to the drying step during the sample preparation of the previous studies. Because the physiological and physicochemical properties of the cell surface would be significantly altered during/after drying,37 the bonds between the bacterium and substratum might be 11550
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir
ARTICLE
Figure 6. (A) Height image of an adherent E. coli K-12 cell on polished stainless steel scanned by tapping-mode AFM in 100 mM PBS. (B) Height information at the cross section in image A. The red arrow indicates the adherent cell, and white arrows indicate EPS surrounding a bacterial cell.
Figure 7. Lateral detachment forces of (A) LPS, (B) type 1 fimbriae, and (C) CA E. coli strains correlated with the surface rms roughness of six substrata.
changed accordingly and then the adhesion strength might be increased significantly.40 Moreover, the viability of bacteria might also be seriously influenced by the drying process because bacterial cells would die quickly (within 1 h) during solution evaporation.41 In the present study, the sample preparation and measurements were conducted in the liquids (including LB medium and 100 mM PBS) to maintain the bacterial viability and conduct in situ AFM measurements. 4.3. Roles of LPS, Type 1 Fimbria, and CA in the Bacterial Attachment Process. The selection of the three E. coli mutants (i.e., D21f2, D21, and JM109) provided an opportunity to evaluate the roles of the LPS molecular structures during the single-cell attachment. The adhesion experiments in the present study revealed that the O-antigen, rather than other LPS structures (core polysaccharide and KDO), could enhance the phase I and II attachment as well as the transformation ratio (Figure 3 and Table 3). For phase I attachment, although the full LPS molecules on the membrane could lead to more steric repulsion,42 the existence of steric interactions did not prevent the bacterial attachment. Also, the O-antigen of LPS was proven to increase the bacterial attachment to various substrata significantly.10,13,24 This phenomenon might be explained by two hypotheses. First, the O-antigen could form hydrogen bonds with the hydroxyl groups on substrata surfaces or interact with water bound to these surfaces.43 This could significantly increase the bacterial attachment. Second, the cells that contained O-antigen had the least negative charge (Table 1) because the relative large O-antigen could hold charged functional groups that could influence the electrostatic double-layer interactions.10 These features of the O-antigen could increase the adhesion strength in the phase I attachment and provide a relatively stable condition under which to facilitate adherent bacteria transfer into phase II attachment. This is consistent with the results of the transformation ratio of the three mutants in the present study. For phase II attachment, AFM tests showed that JM109 with a full LPS structure had the largest adhesion strength compared to that of the other two mutants. This indicated that hydrogen bonds between the O-antigen and the substrata surface still played significant roles during phase II attachment, especially considering the large number (106 per cell) and high coverage (about 75%) of LPS molecules on the bacterial outer membrane.44 Type 1 fimbriae, also known as type 1 pili, are proteinaceous appendages on the surfaces of many gram-negative and grampositive bacteria. Previous studies showed contradictory results when considering the role of type 1 fimbriae in bacterial initial attachment. Several studies suggested that the type 1 fimbriae 11551
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir were critical to the initial bacterial attachment because the attachment of bacteria to type 1 fimbriae significantly exceeded that of those without type 1 fimbriae when mutants attached to various inert surfaces.2,45,46 However, a few other studies revealed that the type 1 fimbriae were not required for the initial bacterial adhesion.33,47 The discrepancy in previous studies might be mainly due to the differences in applied mutants and substrata, cultivation environments, quantification methods, and the attachment phases. In the present study, the results demonstrated that the type 1 fimbriae could largely increase the phase I attachment and the transformation ratio between phases I and II (Figure 4A and Table 3). The hydrophobicity and surface charge tests indicated that the presence of type 1 fimbriae on the cell membrane could slightly change the bacterial surface physicochemical properties (Table 1). However, these changes could not interpret such a large enhancement (Figure 4A) in the phase I attachment. Thus, other fimbrial features might contribute to the initial attachment. Previous studies suggested that type 1 fimbriae could adhere to eukaryotic cell surfaces through the specific recognition of mannoside-containing receptors.33,46 However, mannose was absence in the medium and solutions in the present study, and the presence of type 1 fimbriae still largely enhanced the bacterial phase I attachment to inert surfaces. This might indicate that the type 1 fimbriae, besides binding eukaryotic mannose receptors, could nonspecifically adhere to inert surfaces during the bacterial phase I attachment.45 For phase II attachment, the AFM experiments presenting the type 1 fimbriae did not play important roles in phase II attachment because the measured lateral detachment forces were not significantly different between E. coli K-12 and type 1 fimbriae defective mutant JW4277 (Figure 4B), indicating that the adhesion strength provided by the fimbrial adhesion contributed less, compared with EPS that were excreted after phase I attachment. This was probably caused by the limiting type 1 fimbriae number of one E. coli cell (from 20 to 200 fimbriae per cell, both were counted by TEM48,49), thereby limiting the contribution to the total adhesion strength. CA is a protective capsular polysaccharide surrounding the bacterial cell surface and consists of glucose, galactose, fructose, and glucuronic acid.50 It is reported that CA is synthesized in a biofilm instead of in suspended cells.51 However, a previous study showed that CA could influence the initial interaction between bacteria and substrata by using CA overproduction and defective E. coli C97 mutants.11 In this study, the results using fluorescence microscopy and AFM revealed that CA did not increase the total adherent number of cells and the adhesion strength but rather blocked them in phase I and II attachment, respectively. This is consistent with others’ research results in which CA inhibited bacterial adhesion.11,52 For phase I attachment, variance in the hydrophobicity of each mutant, as a result of different CA contents, might be a possible crucial factor because hydrophobic bacteria adhere to a greater extent than hydrophilic bacteria.53 In this study, the hydrophobicity measurement showed that the CA-overproducing mutant had a more hydrophilic surface than did the defective one by measuring the contact angles of three liquids (Table 1), thus blocking the phase I attachment significantly. For phase II attachment, the existence of capsular CA might block the production of other EPS or contact between substrata surfaces and excreted EPS, thereby decreasing the adhesion strength significantly. This indicated that E. coli avoided excreting CA in the initial attachment and then expressed CA during mature biofilm development11 because
ARTICLE
CA was not directly involved in the phase I and II attachment but rather in the development of biofilm architecture and other phases later by comparing wild-type strains and their mutants.14,15
’ ASSOCIATED CONTENT
bS
Supporting Information. Cluster analysis and canonical correspondence analysis (CCA). Contact angles and surface tensions of applied E. coli strains and substrata. Cluster analysis based on Morisita’s distance for 42 bacteria/substrata combinations (7 E. coli strains 6 substrata) in the phase I attachment. Biplot of CCA for the bacterial attachment process on inert surfaces. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*Phone: (852) 28598551. Fax: (852) 25595337. E-mail: zhangt@ hkucc.hku.hk.
’ ACKNOWLEDGMENT We thank the Hong Kong UGC One-Off Special Equipment Grant Scheme (SEG HKU10) for the financial support on this study. Y.C. thanks the HKU for a postgraduate studentship. The technical assistance of Ms. Vicky Fung is greatly appreciated. ’ REFERENCES (1) Palmer, J.; Flint, S.; Brooks, J. J. Ind. Microbiol. Biotechnol. 2007, 34, 577. (2) Van Houdt, R.; Michiels, C. W. Res. Microbiol. 2005, 156, 626. (3) Zhang, T.; Chao, Y.; Shih, K.; Li, X. Y.; Fang, H. H. P. Ultramicroscopy 2011, 111, 131. (4) Kumar, C. G.; Anand, S. K. Int. J. Food Microbiol. 1998, 42, 9. (5) Carpentier, B.; Cerf, O. J. Appl. Bacteriol. 1993, 75, 499. (6) Van Loosdrecht, M. C. M.; Lyklema, J.; Norde, W.; Schraa, G.; Zehnder, A. J. B. Appl. Environ. Microbiol. 1987, 53, 1898. (7) Dunne, M. W. Clin. Microbiol. Rev. 2002, 15, 155. (8) Burks, G. A.; Velegol, S. B.; Paramonova, E.; Lindenmuth, B. E.; Feick, J. D.; Logan, B. E. Langmuir 2003, 19, 2366. (9) Liu, Y.; Yang, C.; Li, J. Environ. Sci. Technol. 2007, 41, 198. (10) Walker, S. L.; Redman, J. A.; Elimelech, M. Langmuir 2004, 20, 7736. (11) Hanna, A.; Berg, M.; Stout, V.; Razatos, A. Appl. Environ. Microbiol. 2003, 69, 4474. (12) Korea, C.; Badouraly, R.; Prevost, M.; Ghigo, J.; Beloin, C. Environ. Microbiol. 2010, 12, 1957. (13) Li, B.; Logan, B. E. Colloids Surf., B 2004, 36, 81. (14) Balestrino, D.; Ghigo, J.; Charbonnel, N.; Haagensen, J. A. J.; Forestier, C. Environ. Microbiol. 2008, 10, 685. (15) Prigent-Combaret, C.; Prensier, G.; Le Thi, T. T.; Vidal, O.; Philippe, L.; Dorel, C. Environ. Microbiol. 2000, 2, 450. (16) Rose, S. F.; Okere, S.; Hanlon, G. W.; Lloyd, A. W.; Lewis, A. L. J. Mater. Sci.: Mater. Med. 2005, 16, 1003. (17) Senechal, A.; Carrigan, S. D.; Tabrizian, M. Langmuir 2004, 20, 4172. (18) Dickinson, R. B.; Cooper, S. L. Bioeng. Food Nat. Prod. 1995, 41, 2160. (19) Bowen, W. R.; Fenton, A. S.; Lovitt, R. W.; Wright, C. J. Biotechnol. Bioeng. 2002, 79, 170. (20) Hansen, W. R.; Tulyathan, O.; Dawson, S. C.; Cande, W. Z.; Fletcher, D. A. Eukaryotic Cell 2006, 5, 781. (21) Gomez-Suarez, C.; Busscher, H. J.; van der Mei, H. C. Appl. Environ. Microbiol. 2001, 67, 2531. 11552
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
Langmuir
ARTICLE
(22) Deupree, S. M.; Schoenfisch, M. H. Langmuir 2008, 24, 4700. (23) Gaboriaud, F.; Dufr^ene, Y. F. Colloids Surf., B 2007, 54, 10. (24) Li, B.; Logan, B. E. Colloids Surf., B 2005, 41, 153. (25) Vadillo-Rodríguez, V.; Logan, B. E. Environ. Sci. Technol. 2006, 40, 2983. (26) Feil, H.; Feil, W. S.; Detter, J. C.; Purcell, A. H.; Lindow, S. E. Phytopathology 2003, 93, 675. (27) Bergmaier, D.; Lacroix, C.; Macedo, M. G.; Champagne, C. P. Appl. Microbiol. Biotechnol. 2001, 57, 401. (28) Boulos, L.; Prevost, M.; Barbeau, B.; Coallier, J.; Desjardins, R. J. Microbiol. Methods 1999, 37, 77. (29) Brant, J. A.; Childress, A. E. J. Membr. Sci. 2002, 203, 257. (30) Van Oss, C. J. Interfacial Forces in Aqueous Media; Taylor & Francis: Boca Raton, FL, 2006. (31) De Kerchove, A. J.; Elimelech, M. Langmuir 2005, 21, 6462. (32) Atabek, A.; Camesano, T. A. J. Bacteriol. 2007, 189, 8503. (33) Rodrigues, D. F.; Elimelech, M. Biofouling 2009, 25, 401. (34) Razatos, A.; Ong, Y.; Sharma, M. M.; Georgiou, G. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 11059. (35) Sheng, X.; Ting, Y. P.; Pehkonen, S. O. J. Colloid Interface Sci. 2007, 310, 661. (36) Boyd, R. D.; Verran, J.; Jones, M. V.; Bhakoo, M. Langmuir 2002, 18, 2343. (37) Whitehead, K. A.; Rogers, D.; Colligon, J.; Wright, C.; Verran, J. Colloids Surf., B 2006, 51, 44. (38) Truong, V. K.; Lapovok, R.; Estrin, Y. S.; Rundell, S.; Wang, J. Y.; Fluke, C. J.; Crawford, R. J.; Ivanova, E. O. Biomaterials 2010, 31, 3674. (39) He, T.; Shi, Z. L.; Fang, N.; Neoh, K. G.; Kang, E. T.; Chan, V. Biomaterials 2009, 30, 317. (40) Roosjen, A.; Boks, N. P.; van der Mei, H. C.; Busscher, H. J.; Norde, W. Colloids Surf., B 2005, 46, 1. (41) Xie, X.; Li, Y.; Zhang, T.; Fang, H. H. P. Appl. Microbiol. Biotechnol. 2006, 73, 703. (42) Abu-Lail, N. I.; Camesano, T. A. Environ. Sci. Technol. 2003, 37, 2173. (43) Jucker, B. A.; Harms, H.; Hug, S. J.; Zehnder, A. J. B. Colloids Surf., B 1997, 9, 331. (44) Caroff, M.; Karibian, D. Carbohydr. Res. 2003, 338, 2431. (45) Pratt, L. A.; Kolter, R. Mol. Microbiol. 1998, 30, 285. (46) Beloin, C.; Roux, A.; Ghigo, J. M. Curr. Top. Microbiol. Immunol. 2008, 322, 249. (47) Otto, K.; Elwing, H.; Hermansson, M. Colloids Surf., B 1999, 15, 99. (48) Ronald, L. S.; Yakovenko, O.; Yazvenko, N.; Chattopadhyay, S.; Aprikian, P.; Thomas, W. E.; Sokurenko, E. V. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 10937. (49) Sokurenko, E. V.; Courtney, H. S.; Maslow, J.; Siitonen, A.; Hasty, D. L. J. Bacteriol. 1995, 177, 3680. (50) R€att€o, M.; Verhoef, R.; Suihko, M. L.; Blanco, A.; Schols, H. A.; Voragen, A. G. J.; Wilting, R.; Siika-aho, M.; Buchert, J. J. Ind. Microbiol. Biotechnol. 2006, 33, 359. (51) Danese, P. N.; Pratt, L. A.; Kolter, R. J. Bacteriol. 2000, 182, 3593. (52) Razatos, A.; Ong, T. L.; Sharma, M. M.; Georgiou, G. J. Biomater. Sci. Polym. 1998, 9, 1361. (53) An, Y. H.; Friedman, R. J. J. Biomed. Mater. Res. 1998, 43, 338.
11553
dx.doi.org/10.1021/la202534p |Langmuir 2011, 27, 11545–11553
