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Applied and Environmental Microbiology, July 2004, p. 4356-4362, Vol. 70, No. 7
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.7.4356-4362.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

SHORT REPORT

Use of Fluorescent Lectin Probes for Analysis of Footprints from Pseudomonas aeruginosa MDC on Hydrophilic and Hydrophobic Glass Substrata

Eduardo Mora Bejarano and René Peter Schneider*

Department of Microbiology, Institute of Biomedical Sciences, Universidade de São Paulo, São Paulo, São Paulo 05508-900, Brazil

Received 3 September 2003/ Accepted 3 February 2004

ABSTRACT

Microbial footprints of Pseudomonas aeruginosa MDC attached for 1 h to clean or silanized glass were analyzed with fluorescently labeled lectin probes. Footprint composition varied, depending on cell physiology and substratum surface chemistry. This suggests that substratum physicochemistry affected the structure of cell surfaces of adsorbed organisms.

Microbial colonization of surfaces, which may or may not result in the formation of bacterial biofilms, always begins with the adhesion of cells to substrata. Despite the importance of attachment of bacteria to inanimate surfaces in nature, disease, and the deterioration of engineered surfaces, very little is known about the polymers that mediate adhesion of microbes to these surfaces. Putative adhesive biopolymers have been isolated and characterized from the surfaces of planktonic cells (1, 5, 6, 24, 26, 27), an environment very different from the bacterium-substratum interface. Mutants with altered adhesion properties may be used to identify genes of adhesive biopolymers (7, 10, 13, 15, 18, 32), but one has to consider that any mutation that affects cell surface structure is likely to influence adhesion, even if it does not target a polymer directly involved in the establishment of adhesive contacts. Elucidation of the exact role of the polymers identified in these studies in microbial adhesion therefore depends on the development of methods for the direct analysis of the chemical composition of the bacterium-substratum interface.

Microbial footprints are microbial structures left on substratum surfaces after removal of attached cells either by sonication (21, 22) or protease treatment (25) or spontaneous detachment by the shear forces exerted by the liquid on the surface (11). Footprints contain cell surface fragments and excreted materials, such as biosurfactants and other extracellular polymers (11). Footprint formation was first observed by Marshall et al. (19) and has since been reported sporadically by various researchers (11, 17, 25, 31). In the first systematic study on footprint formation, Neu and Marshall (22) revealed the presence of sugar molecules in the footprints of Psychrobacter immobilis SW8. Recently, footprints of Pseudomonas aeruginosa have been analyzed using X-ray photoelectron microscopy (11).

Lectins have been employed by a variety of researchers to analyze sugar composition of biofilm macromolecules (3, 23) and P. aeruginosa biofilms (30, 33). Neu and Marshall (22) used an indirect lectin labeling method in their footprint studies with the marine organism Psychrobacter immobilis SW8. Lectin binding was detected after overnight incubation of samples with nonspecific fluorescent protein probes. In this work, direct lectin assays with fluorescently labeled molecules and fluorescent protein probes were employed to probe the chemical composition of microbial footprints from P. aeruginosa MDC, which was isolated from a water tank at a factory in the outskirts of the city of São Paulo, Brazil, and identified to the species level using biochemical tests and by partial 16S ribosomal DNA sequencing.

Chemicals were reagent grade or better, except where stated otherwise. Glassware was washed with a detergent (neutral MA-02 extran detergent [2% in distilled water]; Merck, São Paulo, Brazil) and then with tap water and rinsed with distilled water. After the glassware had dried, it was soaked in chromic acid solution (2.6%) for 24 h to eliminate organic residues, rinsed in distilled water, and sterilized in an autoclave for 20 min at a pressure of 1 bar. The strain was incubated on a rotary shaker at 37°C in Erlenmeyer flasks containing 100 ml of sterile tryptic soy broth. For preservation, 1-ml samples of cultures in mid-log growth phase were transferred to Eppendorf tubes, glycerol was added to a final concentration of 20% (vol/vol), and the tubes were stored frozen at –80°C. For the production of microbial footprints, cells were harvested by centrifugation in a desktop centrifuge at 4°C at 6,000 rpm for 5 min either after they were allowed to grow overnight (stationary phase; total cultivation time, 15 h) or when the optical density at 600 nm reached a value around 0.55 (mid-log phase of growth). The cells were washed twice with a filter-sterilized NaCl solution (0.02 M NaCl, buffered with 5 mM phosphate to pH 7.0). The washed cells were resuspended in the same solution to an optical density at 600 nm of 1.0 and stored at room temperature prior to use.

Hydrophilic glass slides (24 by 60 mm, cut from 0.9-mm-thick borosilicate glass surfaces supplied by Jena Glass, Jena, Germany) that were either clean or made hydrophobic by the silanization method of Neu and Marshall (22) were used as substrata for preparation of footprints. The slides were cleaned by immersion in chromic acid solution overnight. The slides were then washed with distilled water and dried at 50°C. Substrata were placed inside covered petri dishes for drying and storage. Adhesion assays were initiated by placing 1 drop (250 µl) of cell suspension on the substratum surface. After 1 h, nonadhered cells were removed by washing the cells five times with filter-sterilized NaCl solution. Irreversibly attached cells were counted immediately after the washing step, after attached cells were labeled with 4',6'-diamidino-2-phenylindole (DAPI). The average number as well as standard deviation of cells counted on 15 different fields of a substratum were determined and converted into the number of cells per square centimeter. Adhesion experiments were repeated at least three times with independently grown cultures. Comparing the results with a Student t test (P = 0.99) revealed no statistically significant differences in adhesion between replicate experiments.

P. aeruginosa MDC adhered to the substratum surfaces either along the long axis of the cell or via the nonflagellated cell pole. In the latter case, the only contact between the cell and substratum occurred at the polar region. There was no statistically significant difference in the numbers of stationary-phase and exponential-phase cells retained on nonsilanized hydrophilic (10.7 x 104± 474 mid-log-phase cells/cm2 and 11.6 x 104 ± 524 stationary-phase cells/cm2) or silanized hydrophobic substratum surfaces (9.6 x 104 ± 725 mid-log-phase cells/cm2 and 10.4 x 104 ± 1,000 stationary-phase cells/cm2). No statistically relevant variation of the mean values of retained cells was observed in independent adhesion assays performed during a 10-week period with independently grown cells.

Lectins (Table 1) labeled with fluorescein isothiocyanate purchased from Sigma Chemical Company (St. Louis, Mo.) were dissolved (1 mg/ml) in phosphate buffer (15 mM, pH 7.0), filtered through 0.22-µm-pore-size membranes, and stored at 4°C in the dark. Samples (50 µl) of lectin solution were applied to substratum surfaces and incubated in the dark for 4.5 h inside a moistened petri dish. Substrata were washed three times with filter-sterilized NaCl solution prior to fluorescence microscopy with an Axiovert S100 inverted microscope (with 495-nm-wavelength [excitation] and 535-nm-wavelength [emission] filters; Zeiss, Jena, Germany) fitted with a Hamamatsu C5810 digital camera (Sanyo Denki, Tokyo, Japan). Images were analyzed with KS-400 image analysis software. Griffonia simplicifolia lectin 1 (BS-1) did not bind to any cell surface structure. Points of intense fluorescence on cell surfaces surrounded by less intensively fluorescing areas demonstrate that compounds recognized by concanavalin A (ConA) occurred in clusters on the surfaces of exponential- and stationary-phase cells of P. aeruginosa MDC attached to hydrophilic and hydrophobic substrata (Fig. 1 [only exponential-phase cells are shown]). Similar results were obtained with Ulex europeaus lectin I (UEA-I).


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  		TABLE 1. Sugar specificities of FITC-labeled lectins used in this study

 


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  		FIG. 1. Interaction of fluorescently labeled PHA-P (A and B) and ConA (C) lectins with attached exponential-phase cells on nonsilanized hydrophilic glass (A and C) and silanized hydrophobic glass (B). Magnification, x1,000. Bar, 10 µm.

 
The differences in binding of the lectins WGA (wheat germ agglutinin) and PHA-P (phytohemagglutinin from Phaseolus vulgaris) to surfaces of cells attached to silanized and nonsilanized substrata provided evidence for substratum-induced modifications of cell surface organization. Planktonic exponential- and stationary-phase cells incubated with lectins for the same amount of time as adsorbed cells were labeled similarly to cells adsorbed on hydrophilic substrata, which bound the PHA-P lectin nonuniformly (Fig. 1A) in a manner similar to that of the ConA lectin (Fig. 1C). Only a few molecules of the WGA lectin interacted with these cells (not shown). The similarity of labeling in cells suspended in aquatic medium and cells adsorbed on nonsilanized glass probably reflected the hydrophilic nature of these interfaces.

On hydrophobic silanized glass, however, exponential- and stationary-phase cells from the same cell suspensions were labeled uniformly by the PHA-P and WGA lectins (Fig. 1B [only PHA-P shown]). The same results were observed after only 30-min incubation of lectins with cells, suggesting a rapid cell surface rearrangement on hydrophobic substrata probably immediately after contact and subsequent maintenance of the substratum-specific cell surface structure for a period of at least 3.5 h. In this molecular surface rearrangement process, which probably reflects the adaptation to a hydrophobic interface of a cell surface equilibrated in a hydrophilic solution, compounds recognized by PHA-P, which occurred in clusters on planktonic cells, became uniformly distributed over the cell surface. The same phenomenon occurred with molecules that interacted with WGA, which were not exposed on the surfaces of planktonic cells. Clearly, P. aeruginosa MDC cell surfaces react dynamically to the physicochemical conditions of the substratum surface and redistribute cell surface molecules accordingly. Membrane fluidity may assist this substratum-specific cell surface response.

For cell removal, substrata were immersed in glass beakers filled with filter-sterilized NaCl and placed in a sonication bath (Ultrasonik 28x NDI; NEY, Inc., Bloomfield, Conn.). After sonication, the substrata were washed three times with the same buffer solution. All experiments were performed in triplicate. Three minutes of sonication at energy level 10 at 37°C with 10 min of degassing was sufficient for complete removal of attached cells from surfaces and abundant production of footprints (Fig. 2). Sonication did not kill or inactivate desorbed organisms. Plate cell counts (tryptic soy agar plates) and total (DAPI-labeled) cell counts were statistically similar in sonication supernatants after cell removal (Table 2). Phase-contrast microscopy revealed that substratum surface chemistry apparently influenced cell desorption during sonication (Fig. 3). All cells that remained on clean glass surfaces after sonication were attached via one cell pole and protruded at variable angles from the substratum surface (Fig. 3). The adhesive polymers at the contact point were fluorescently labeled by the PHA-P lectin (Fig. 3). The nonfluorescent notch at the center of each microbial footprint shown in Fig. 3 is the shadow of the cell body. These shadows were formed because the substratum was illuminated from below and very little PHA-P lectin adsorbed in regions other than the cell pole. Sonication of cells attached along their long axis on hydrophilic glass apparently caused rupture of cell-substratum bonds on one side first, before further sonication released the cell from the surface. No such cell-pole contacts were observed with the PHA-P lectin on silanized glass.



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  		FIG. 2. P. aeruginosa MDC and footprints on hydrophilic glass surfaces. Exponential-phase cells retained on the substratum after 1 h, surfaces that were not sonicated (A and B), and cells and footprints after 1 min (C and D), 2 min (E and F), and 3 min (G and H) of sonication. The arrows point to footprints. Scale bars: 5 µm (A, C, and E); 2 µm (D); and 1 µm (B, F, G, and H).

 

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  		TABLE 2. Cell survival in sonication buffers after termination of sonication

 


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  		FIG. 3. Interaction of the PHA-P lectin with P. aeruginosa MDC cells. (A) Interaction of the PHA-P lectin from Phaseolus vulgaris with P. aeruginosa MDC cells attached via their polar ends on hydrophilic glass, as analyzed by fluorescence microscopy. Mild sonication conditions that allowed for removal of most, but not all, attached microbes were used. Note labeling of the contact point between the cell and substratum only. Magnification, x1,000. Bar, 10 µm. (B) Schematic representation of the labeling point shown in panel A. The arrows point to cells (identified by phase-contrast imaging of the same microscopic field).

 
For electron microscopy, substrata were fixed with a solution of 8% glutaraldehyde (vol/vol) and 4% Formol (vol/vol) in phosphate buffer (0.1 M, pH 7.0) for 2 h. Samples were subsequently dehydrated in a series of ethanol solutions (30 min [each] in 50%, 70%, and 90% ethanol solutions, followed by four periods of 15 min in 100% ethanol), dried in a critical point dryer (Baltec CPD 030; Balzers, Fürstentum, Lichtenstein), and metallized with a thin gold film approximately 15 nm thick (Balzers Union SCP 040 metallizer; Balzers, Fürstentum Lichtenstein) and subsequently analyzed by scanning electron microscopy using a Philips XL-30 electron microscope (Windhoven, The Netherlands). Scanning electron micrographs of stationary- or exponential-phase cells attached to either hydrophilic or hydrophobic substrata were similar; only micrographs of exponential-phase cells attached to hydrophilic glass are shown in Fig. 2. Most footprints were not geometric imprints of cells, but structures of irregular morphologies smaller than a cell diameter (Fig. 2). Fibrils visible in Fig. 2B are probably extracellular polymeric substances that condensed after dehydration and demonstrate the possible involvement of excreted polymers in microbial footprints.

Lectin binding to microbial footprints was investigated with substrata subjected to 2 min of sonication at energy level 7 and 30°C bath temperature to remove most, but not all, cells in order to directly compare the fluorescence signals of attached cells and footprints in the same microscopic field. The positions of the few remaining attached cells in a particular field were determined by phase-contrast microscopy after fluorescence analysis. Nonspecific adsorption of ConA, PHA-P, and BS-1 occurred on a small scale on both hydrophilic and hydrophobic glass surfaces as evidenced by a faint, homogeneous fluorescence that appeared after incubation of the clean substrata with lectin solutions. UEA-I did absorb nonspecifically only to hydrophobic substrata, and WGA did not adsorb to clean hydrophilic and hydrophobic surfaces.

Lectin-labeled microbial footprints were circular or elliptic dots with surface areas much smaller than the contact area between cells and substrata, a shape similar to that observed in scanning electron micrographs (Fig. 4 [only ConA shown]). The shape of footprints depends on the number of contact points between the cell surface and substratum. Cells attached via one of their cell poles will produce dot-shaped microbial footprints. The shape of microbial footprints from cells attached along the long axis of the cell may vary anywhere from a dot pattern to full copies of the contact zone between the cell and substratum, as demonstrated by Neu and Marshall (22), depending on the number and location of contact points with the substratum. Fibrils were detected with PHA-P and WGA in microbial footprints on nonsilanized glass (Fig. 5), but not on silanized glass (not shown). These fibrils most probably were present on the surfaces of cells attached to both substrata but formed relatively strong contacts only on nonsilanized glass, which caused their separation from the cell surface during sonication. The background fluorescence of WGA on nonsilanized glass in Fig. 5 is evidence for adsorption of a conditioning film of molecules containing sugars on this surface, which had been in contact with microbes.



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  		FIG. 4. Interaction of the ConA lectin on hydrophilic glass with cells and footprints of P. aeruginosa MDC as analyzed by fluorescence microscopy. Mild sonication conditions that allowed for removal of most, but not all attached, microbes were used. Footprints and only a few cells from exponential-phase cultures (A) and stationary-phase cultures (B) are shown. The arrows point to cells (identified by phase-contrast imaging of the same microscopic field). The numbers of cells attached to both substrata were similar prior to sonication. Note the much stronger fluorescence signal on the surface covered with stationary-phase cells. Magnification, x1,000. Bar, 10 µm.

 


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  		FIG. 5. Fibrils on hydrophilic glass detected with lectins PHA-P (A) and WGA (B). The faint background signal is due to nonspecific adsorption of the lectins in small quantities to the substratum surface for PHA-P. For WGA, the faint background signal indicates adsorption of a conditioning film with sugar-containing molecules, since no such background signal was observed with this lectin on clean surfaces. The arrows point to cells (identified by phase-contrast imaging of the same microscopic field). Magnification, x1,000. Bar, 10 µm.

 
Although a similar number of stationary- and exponential-phase cells attached to substrata, microbial footprints produced by stationary-phase cells contained a larger number of binding sites for ConA than microbial footprints from exponential-phase cells, as evidenced by the more intense fluorescence signal observed on substrata covered with material from stationary-phase cells (Fig. 4). The intensities of fluorescence emitted from individual microbial footprint structures were similar on both surfaces. Similar results were obtained with UEA-I, PHA-P, and WGA. Clearly, more cell surface polymers were able to form strong links with the substratum in stationary-phase cells.

Changes in the expression of band A and band B lipopolysaccharides (LPS) in exponential- and mid-log-phase cells may explain the differences in footprint composition in stationary- and log-phase cells. LPS are nearly linear molecules with a length of between 35 to 40 nm in the case of P. aeruginosa (2). Long-chain band B LPS do mask shorter cell surface structures, such as band A LPS or surface proteins (20). The relative amounts of long- and short-chain LPS on the surface of P. aeruginosa are physiological variables and depend on the growth conditions of the organism (16, 18, 20). If long-chain LPS had dominated the cell surfaces of exponential-phase organisms, these molecules would have limited the contact of shorter cell surface structures with the substratum, resulting in detection of less protein and in the observation of fainter lectin signals. Reduced amounts of long-chain band B LPS in stationary-phase cells would allow more cell surface structures, such as proteins or short-chain band A LPS, to contact the substratum surface. In this instance, the mass of cell surface structures contained in footprints would be higher than in microbial footprints from exponential-phase cells.

An alternative explanation for the differences in microbial footprint composition in exponential- and stationary-phase cells is a shift in the plane of the shear forces inside the cell surface of the two types of organisms. Microbial footprints contain the adhesive biopolymers and associated cell surface polymers whose binding force to the substratum surface is stronger than the cohesive forces, which retain them in the cell surface. A shift in the plane of the shear forces due to alterations in the composition of cell surface structures in mid-log-phase and stationary-phase cells could also account for some of the changes in footprint composition. The denser microbial footprint coverage of surfaces exposed to stationary-phase cells may also reflect the production of a larger quantity of extracellular polysaccharides (EPS) by this phenotype, which were deposited together with the cells on substratum surfaces.

One hundred fifty microliters of each lectin solution was preincubated individually with 150 µl of each sugar (Table 1) at concentrations of 50 and 100 mg/ml (stock solutions prepared by dissolving sugars in the same buffer used for lectin preparation) in the dark for 30 min prior to application of a 150-µl volume of the lectin-sugar mixture on substratum surfaces. Preincubation of lectins with sugars totally suppressed lectin binding to microbial footprints, thus demonstrating the specific nature of the interaction. Lectin binding to footprints and cell surfaces does not permit a priori identification of the type of biopolymer present in the structure. It means simply that the type of sugar linkage the lectin recognizes is present in the sample.

Several macromolecules known to be located on the surface of P. aeruginosa contain sugars recognized by the lectins employed in this study. LPS, a molecule known to be released spontaneously into the medium during growth of P. aeruginosa (4), contains glucose, mannose, and N-acetylglucosamine, sugars recognized by ConA and PHA-P, as well as N-acetylfucosamine, recognized by the lectin UEA-I (14). Uronic acid residues recognized by WGA are present in alginate, an EPS produced abundantly by mucoid P. aeruginosa when growing in biofilms (33) and in band B LPS (20). However, alginate is not present in the EPS of nonmucoid P. aeruginosa PAO1 and PA14 (34). Cellulose, a glucose polymer potentially recognized by ConA, was reported as a major component of Pseudomonas fluorescens EPS (28).

Several proteins located on the surface of P. aeruginosa are glycoproteins that contain sugar residues recognized by the lectins used in this study. For example, pilin contains the sugar FucNAc recognized by UEA-I (7). Sugar-containing cell surface molecules have been implicated in the adhesion of P. aeruginosa to solid substrata (1, 9, 18, 29, 33). Rhamnolipids, an important group of compounds involved in biofilm formation (8) that contain fatty acids linked to rhamnose sugars (12) would not have been detected by the lectins employed in this work.

ACKNOWLEDGMENTS

This work was supported in part by FAPESP grants 95/09511-8 and CNPQ grant 30.0730/00-4 to R.P.S. and FAPESP MSc Fellowship 00/11550-1 to E.M.B.


   FOOTNOTES
 
* Corresponding author. Mailing address: Departamento de Microbiologia, Instituto de Ciências Biomédicas, Universidade de São Paulo, Av. Prof. Lineu Prestes, 1374, CEP 05508-900, São Paulo, São Paulo, Brazil. Phone: 55 11 3091 7745. Fax: 55 11 3091 7354. E-mail: schneide{at}icb.usp.br. Back

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Applied and Environmental Microbiology, July 2004, p. 4356-4362, Vol. 70, No. 7
0099-2240/04/$08.00+0     DOI: 10.1128/AEM.70.7.4356-4362.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.




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J. Bacteriol. Microbiol. Mol. Biol. Rev. Eukaryot. Cell All ASM Journals