January 2017
Volume 58, Issue 1
Open Access
Cornea  |   January 2017
Bacterial Coaggregation Among the Most Commonly Isolated Bacteria From Contact Lens Cases
Author Affiliations & Notes
  • Ananya Datta
    School of Optometry and Vision Science, University of New South Wales, Sydney, New South Wales, Australia
  • Fiona Stapleton
    School of Optometry and Vision Science, University of New South Wales, Sydney, New South Wales, Australia
  • Mark D. P. Willcox
    School of Optometry and Vision Science, University of New South Wales, Sydney, New South Wales, Australia
  • Correspondence: Mark D. P. Willcox, School of Optometry and Vision Science, University of New South Wales, Sydney, NSW 2052, Australia; m.willcox@unsw.edu.au
Investigative Ophthalmology & Visual Science January 2017, Vol.58, 50-58. doi:10.1167/iovs.16-20593
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      Ananya Datta, Fiona Stapleton, Mark D. P. Willcox; Bacterial Coaggregation Among the Most Commonly Isolated Bacteria From Contact Lens Cases. Invest. Ophthalmol. Vis. Sci. 2017;58(1):50-58. doi: 10.1167/iovs.16-20593.

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      © 2017 Association for Research in Vision and Ophthalmology.

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Abstract

Purpose: To examine the coaggregation and cohesion between the commonly isolated bacteria from contact lens cases.

Methods: Four or five strains each of commonly isolated bacteria from contact lens cases, Staphylococcus aureus, Staphylococcus epidermidis, Pseudomonas aeruginosa, and Serratia marcescens, were grown, washed, mixed in equal proportions, and allowed to coaggregate for 24 hours. Lactose (0.06 M), sucrose (0.06 M), and pronase (2 mg/mL; 2 hours, 37°C) were used to inhibit coaggregation. Oral bacterial isolates of Actinomyces naeslundii and Streptococcus sanguinis were used as a positive control for coaggregation. Cohesion was performed with the ocular bacteria that demonstrated the highest level of coaggregation. Production of growth-inhibitory substances was measured by growing strains together on agar plates.

Results: The oral bacterial pair showed >80% coaggregation. Coaggregation occurred between ocular strains of S. aureus (2/5) or S. epidermidis (2/5) with P. aeruginosa strains (3/5); 42% to 62%. There was only slight coaggregation between staphylococci and S. marcescens. Staphylococcus aureus coaggregated with S. epidermidis. Lactose or sucrose treatment of S. aureus but pronase treatment of P. aeruginosa reversed the coaggregation. There was no cohesion between the ocular isolates. P. aeruginosa was able to stop growth of S. aureus but not vice versa.

Conclusions: This study demonstrated for the first time that ocular isolates of P. aeruginosa and S. aureus could coaggregate, probably through lectin–carbohydrate interactions. However, this may not be related to biofilm formation in contact lens cases, as there was no evidence that the coaggregation was associated with cohesion between the strains.

Microbial contamination is frequently observed during the use of medical devices. Contact lenses and their accessories such as contact lens cases can become contaminated by microbes.15 Contact lens cases have been reported to be more prone to biofilm formation than contact lenses.5,6 Adhesion and subsequent biofilm formation by the microbes on contact lenses or cases may lead to the development of keratitis.7 
The frequency of microbial contamination of contact lens storage cases ranges from 30% to 85% during asymptomatic contact lens wear despite the use of contact lens disinfecting solutions.1,2,5,8,9 Contamination rate and types of microbes can vary between different studies depending on factors such as the sample population, bacterial culture conditions, and inclusion of symptomatic or asymptomatic wearers or those with microbial keratitis.2,1013 A variety of micro-organisms have been isolated and identified in contact lens storage cases including many types of bacteria, fungi, and protozoa.6,8,14 Bacterial contamination is the most common and can occur in approximately 77% of lens cases, followed by fungi (24%) and protozoa (20%).8 The most commonly found bacteria from lens cases are Staphylococcus epidermidis and Staphylococcus aureus.2,5 Staphylococcus epidermidis is commonly found as a part of the normal external ocular microbiota,15,16 and S. aureus is the second most commonly isolated microbe from contact lens cases of both asymptomatic contact lens wearers and those with microbial keratitis.2,10,12 Among Gram-negative bacteria, Pseudomonas aeruginosa and Serratia spp are frequently isolated from contact lens cases2,5,10 and are well-known opportunistic pathogens involved in microbial keratitis.17,18 
Bacterial biofilm formation involves several stages including adhesion, microcolony formation, maturation, and eventual dispersal. Within the process of adhesion and bacterial microcolony formation, microbial coaggregation and cohesion may occur and contribute to biofilm formation and maturation. Coaggregation is the specific recognition and adherence of bacteria to themselves or other microbial types and can be associated with the development of complex multispecies biofilm.1921 Coaggregation was first observed between bacteria isolated from human dental plaque22,23 and has been observed in bacterial strains isolated from other places including fresh water.20 Coaggregation is often mediated by protein–saccharide interactions and can be blocked by the addition of simple sugars. The bacterial proteins and saccharides can be found on flagella and pili.2428 Coaggregation can occur with bacteria in suspension or when one type is bound to a surface. Cohesion is the process whereby the adhesion of one type of microbe promotes the subsequent adhesion of other microbes. This can occur by pioneer microbes providing new surfaces (their own surface) for other bacteria to adhere to (i.e., via coaggregation) or by the pioneer microbes modifying the surface itself to facilitate adhesion of secondary colonizers. There have been no reports of the cohesion abilities and the mechanism of interaction of bacteria isolated from contact lens cases. 
The aim of this present study was to examine the coaggregation pattern, mechanism of interaction, and cohesion between the most commonly isolated bacteria from contact lens cases, P. aeruginosa, Serratia marcescens, S. aureus, and S. epidermidis. Knowledge of these interactions may aid in the rational design of strategies to reduce adhesion and biofilm formation by bacteria in contact lens cases. 
Methods
Microbial Strains
Bacterial strains and their sources are listed in Table 1. Five strains each of P. aeruginosa, S. marcescens, and S. aureus and four strains of S. epidermidis were used in this study. Actinomyces naeslundii and Streptococcus sanguinis (Table 1) isolated from dental plaque were used as positive controls for coaggregation.23 
Table 1
 
Bacteria Used in the Study
Table 1
 
Bacteria Used in the Study
Coaggregation Assay
Gram-negative bacteria were grown in Luria broth (LB; Sigma-Aldrich, Castle Hill, NSW, Australia) and Gram-positive bacteria in brain heart infusion (BHI: Becton, Dickinson, Macquarie Park, NSW, Australia) for 18 to 24 hours at 37°C. Bacterial cells were collected by centrifugation at 10,000g for 15 minutes at 4°C and washed three times in coaggregation buffer (1 mM Tris (hydroxymethy) amino methane, 0.1 mM CaCl2, 0.1 mM MgCl2, 0.15 M NaCl, and 3.1 mM NaN3 at pH 8.0). After the cells were washed, the optical density (OD) of the bacterial strains was adjusted to 1.0 (1 × 109 CFU mL−1) at 660 nm using a spectrophotometer (FLUOstar Omega; BMG Labtech, Ortenberg, Germany). An equal volume (0.2 mL) of two cell suspensions was mixed (e.g., P. aeruginosa 6294 and S. aureus 31) and vortexed for 30 seconds, and the OD was measured immediately at 660 nm (OD1). Then the cell suspension was centrifuged for 2 minutes at 650g and left to stand at ambient temperature for 2 hours, and the OD660 of 0.2 mL of the upper layer was measured (OD2).29 This procedure was repeated again after 24 hours (OD24). The percentage coaggregation was assessed using the following formula:    
The percentage decrease in OD was determined and mean and standard deviation were calculated. The experiments were repeated four times. A percentage decrease greater than 30% indicated coaggregation.30 
Inhibition of Bacterial Coaggregation
Only those coaggregating pairs that showed >30% coaggregation were tested using inhibitor substances. Lactose and sucrose were used to reverse the coaggregation reactions as described by Cisar et al. (1979).23 Lactose (0.06 M) or sucrose (0.06 M) was prepared in coaggregation buffer and was individually added to one of the bacterial aggregating pairs.31 Individual bacteria from the coaggregation pairs were also treated with protease (pronase from Streptomyces griseus, P-5130; Sigma-Aldrich) at a final enzyme concentration of 2 mg/mL. The bacterial strains were incubated for 2 hours in lactose, sucrose, or pronase at ambient temperature and then the cell suspensions were washed three times with coaggregation buffer to remove unbound inhibitors. The lactose-, sucrose-, or pronase-treated bacteria were added with the nontreated bacteria partners, and the percentage of coaggregation was recorded after 2 hours of incubation at ambient temperature as described previously. Inhibition of coaggregation was measured by comparing the coaggregation in the absence or presence of the sugars or pronase.21,23 A solution of 0.05% Tween-20, 0.2 M NaCl was used as a negative control to control for nonspecific bacterial interactions such as those associated with hydrophobicity and ionicity. Only if there was a greater inhibition than this control was inhibition considered to have occurred. 
Additionally, the ability of lectin-specific sugars (D-galactose and L-fucose) to inhibit coaggregation of P. aeruginosa was assessed. Pseudomonas aeruginosa lectins LecA (PA-1) and LecB (PA-II) bind specifically with D-galactose and L-fucose, respectively.32 In this assay, P. aeruginosa was incubated in coaggregation buffer along with D-galactose (0.05 M) or L-fucose (0.05 M) at pH 8.0. After washing and incubating with nontreated cells, the coaggregation was compared as described above. 
Bacterial Motility Test
A twitching motility assay was performed as described by O'Toole and Roberto (1998).28 Bacteria were grown as described for the coaggregation assay, washed, and resuspended in phosphate-buffered saline (PBS; pH 7.4 NaCl 8 g 1−1, KCl 0.2 g 1−1, Na2HPO4 1.15 g 1−1, KH2PO4 0.2 g 1−1 pH 7.2) at an OD of 1.0 at 660 nm. The cells were then stab-inoculated into agar plates (1% [wt/vol] tryptone; 0.5% [wt/vol] purified agar; Oxoid, Basingstoke, UK); 0.5% [wt/vol] NaCl; ICN Biomedicals, Irvine, CA, USA) using a sterile stainless steel stick and incubated overnight at 37°C. After incubation the plates were kept at ambient temperature for 2 days. Twitching motility was observed as a faint turbid zone around the stab. The zone and pattern of motility were examined at the bottom of the agar plate. The twitching and swarming motility were differentiated on agar plate by the pattern of their growth. Swarming motility was distinguished from swimming by the appearance of dendritic patterns in the bacterial growth that elongated and branched from a central colony. 
Additionally, flagella-mediated swimming motility of bacteria was examined using the hanging drop technique. One drop of sterile PBS was placed on the center of a glass slide, and a loopful of bacterial suspension grown as previously described was added. A sterile coverslip was placed over the suspension and the coverslip/glass slide edges were sealed to maintain humidity. Flagella-mediated motility was observed in the drop using a light microscope (Leitz Wetzlar Microscope, Wetzlar, Germany) at ×200 magnification, and care was taken to differentiate active motility from random Brownian motion. The procedure was repeated three times. 
Cohesion Assay
The cohesion assay was performed using only the bacterial pairs that showed the highest coaggregation. Strains were grown and adjusted to 0.1 OD660 as described for the coaggregation assay. Pseudomonas aeruginosa 6294 (2 mL) was added to contact lens storage cases (Re-Nu Multiplus; Bausch & Lomb, Rochester, NY, USA), and the case was loosely recapped and incubated at 37°C for 24 hours. The lens cases were washed three times with PBS to remove loosely adherent cells, and 2 mL S. aureus 31 suspension was then added and incubated at 37°C for 24 hours. Alternatively, the lens cases were first incubated with S. aureus 31 and then with P. aeruginosa 6294. 
After overnight incubation with the second type of bacterium, lens cases were rinsed gently twice with PBS to remove loosely attached cells. Subsequently, 2 mL PBS was added to each well of the lens case along with a sterile magnetic stirring bar, and the case was vortexed for 1 minute to dislodge the adherent bacterial cells. Tenfold serial dilutions of the dislodged bacteria were made in PBS, and the dilutions were plated on selective isolation agar Cetrimide (Thermo Fisher Scientific, Scoresby, VIC, for P. aeruginosa and Mannitol salt agar (Thermo Fisher Scientific) for S. aureus. Plates were incubated for 18 to 24 hours at 37°C to determine the number of viable bacteria. Controls used a single type of bacterial cell incubated for 24 hours at 37°C followed by addition 2 mL sterile PBS and incubation for a further 24 hours at 37°C and processed as described. The experiments were repeated on three different occasions with three samples each time. 
Inhibition of Bacterial Growth
This assay was performed only on strains used in the cohesion assay. In the first instance, S. aureus 31 was grown overnight in BHI and resuspended to 0.1 OD660 in PBS and spread on a nutrient agar plate (Thermo Fisher Scientific) using a sterile cotton swab. Subsequently, 10 μL P. aeruginosa 6294, grown overnight in LB broth and resuspended to 0.1 OD660 in PBS, was spotted onto the S. aureus lawn and the plate incubated overnight at 37°C. The same procedure was performed by spotting S. aureus culture on a lawn of P. aeruginosa. After incubation, inhibition of growth was assessed and graded as follows: ‘‘No inhibition'' was indicated when no zone of inhibition was observed around the bacterial spot of inoculation; ‘‘weak inhibition'' was indicated by an inhibition halo ≤ 6 mm around the bacterial spot inoculation; ‘‘strong inhibition'' was indicated by an inhibition halo > 6 and ≤ 16 mm; and ‘‘very strong inhibition'' was indicated by an inhibition halo > 16 mm.33 The inhibitory zone was expressed in millimeters from three independent experiments measured directly on the agar plates. 
Statistical Analysis
Data analysis was performed using Microsoft Excel 2010 (Redmond, WA, USA) and Statistical Package for Social Science for Windows version 20.0 (SPSS, Inc., Chicago, IL, USA). The percentage of bacterial coaggregation between different pairs and groups was compared using ANOVA. Additionally, the comparison of the percentage of coaggregation between different time points was performed using paired sample t-test. 
Results
Bacterial coaggregation occurred between different bacterial pairs. In most cases where coaggregation was observed it occurred after 2 hours of incubation and was only slightly increased at 24 hours. Four out of five S. aureus and three out of four S. epidermidis strains were able to coaggregate with some of the P. aeruginosa strains after 24 hours incubation. Pseudomonas aeruginosa 6294 coaggregated with all strains of S. aureus; P. aeruginosa 6206 coaggregated with all S. aureus strains except ATCC 6538; P. aeruginosa Paer1 coaggregated with S. aureus strains 31, 38, and 62; and P. aeruginosa ATCC 9027 only with S. aureus strain 31 (Table 2). The highest percentage coaggregation of 62 ± 3% was observed between P. aeruginosa 6294 and S. aureus 31 after 24 hours incubation. Pseudomonas aeruginosa 149 did not coaggregate with any strain of S. aureus, and P. aeruginosa ATCC 9027 a weak coaggregation with only S. aureus 31 after 24 hours incubation. Pseudomonas aeruginosa (6294 and 6206) showed significant coaggregation (P < 0.001) with S. epidermidis strains ATCC 35984 and NCTC 11047, and the maximum percentage of coaggregation was 49 ± 2% between P. aeruginosa 6294 and S. epidermidis ATCC 35984 after 24 hours. Pseudomonas aeruginosa Paer1 showed significant coaggregation with S. epidermidis NCTC 11047 only. Serratia marcescens (ATCC 13880, 27, 5, and 32) showed low and sporadic coaggregation (>30%; P < 0.05) with S. aureus (31, 38, 61, and 62) after 24 hours incubation, but the percentage of coaggregation did not exceed 33% (Table 2). Only S. marcescens 35 coaggregated with S. epidermidis, giving a low-level (30%) coaggregation with S. epidermidis ATCC 35984 after 2 or 24 hours incubation. There was no coaggregation between the two Gram-negative bacteria P. aeruginosa and S. marcescens (data not shown). However, the two Gram-positive bacteria showed coaggregation. Staphylococcus aureus strains 31 and 38 showed significant coaggregation (P < 0.001) after 24 hours with S. epidermidis strains ATCC 35984 (52 ± 3%; 47 ± 2%, respectively; Fig. 1) and NCTC 11047 (46 ± 3%; 42 ± 2%, respectively; Fig. 1). Staphylococcus epidermidis 19 coaggregated with S. aureus 31 and 62, and S. epidermidis 5 coaggregated with S. aureus 38 after 24 hours. The positive control of three A. naeslundii strains and S. sanguinis CR2B showed 92 ± 3% coaggregation after 24 hours (Fig. 1). 
Table 2
 
Coaggregation Between Gram-Positive and Gram-Negative Bacteria After 2 or 24 Hours of Incubation
Table 2
 
Coaggregation Between Gram-Positive and Gram-Negative Bacteria After 2 or 24 Hours of Incubation
Figure 1
 
Coaggregation between S. aureus (SA) and S. epidermidis (SE) strains. Positive coaggregation was considered to be ≥30% and is indicated by the line on the graph. Also shown is the percentage coaggregation of the positive control strains: Actinomyces naeslundii (AN) with Streptococcus sanguinis (SS).
Figure 1
 
Coaggregation between S. aureus (SA) and S. epidermidis (SE) strains. Positive coaggregation was considered to be ≥30% and is indicated by the line on the graph. Also shown is the percentage coaggregation of the positive control strains: Actinomyces naeslundii (AN) with Streptococcus sanguinis (SS).
The results of the inhibition of coaggregation are shown in Table 3. The negative control solution of Tween-20 and NaCl (used to determine any coaggregation as the result of hydrophobic or charge interactions) gave reductions of coaggregation of between 30% and 38%; therefore a significant inhibition in coaggregation for any treatment was considered to be ≥40%. For the coaggregating pairs of P. aeruginosa with S. aureus or S. epidermidis, treatment of P. aeruginosa with lactose or sucrose did not inhibit coaggregation (<40% inhibition), whereas treatment of S. aureus or S. epidermidis strains with lactose or sucrose inhibited coaggregation (P < 0.05) for the most pairs (the exception being P. aeruginosa Paer1 and S. aureus 38; Table 3). Lactose or sucrose treatment of S. aureus 31 gave 52 ± 3% and 56 ± 4% inhibition of coaggregation with P. aeruginosa 6294. When P. aeruginosa strains were pretreated with either D-galactose or L-fucose there were reductions (P < 0.05) in coaggregation with all tested strains of S. aureus and S. epidermidis (Table 4). Additionally, the treatment of pronase also inhibited coaggregation between certain pairs. The treatment of P. aeruginosa with pronase inhibited coaggregation (P < 0.05) with all the staphylococci, but pronase treatment to staphylococci did not inhibit coaggregation (Table 3). 
Table 3
 
Inhibition of Coaggregation Using Lactose, Sucrose, or Pronase
Table 3
 
Inhibition of Coaggregation Using Lactose, Sucrose, or Pronase
Table 4
 
Inhibition of Coaggregation Using D-Galactose (0.05 M) and L-Fucose (0.05 M)
Table 4
 
Inhibition of Coaggregation Using D-Galactose (0.05 M) and L-Fucose (0.05 M)
While coaggregation was observed between S. aureus and S. epidermidis there was no inhibition of coaggregation when S. aureus strains were treated with either lactose or pronase. However, treatment of S. epidermidis strains with either lactose or pronase inhibited coaggregation with S. aureus. For two out of three pairings, treatment of S. aureus with sucrose inhibited coaggregation with S. epidermidis, but treatment of S. epidermidis with sucrose did not inhibit coaggregation with S. aureus. The coaggregation between S. marcescens and S. aureus was inhibited by pronase treatment but not by treatment with sugars. 
Due to the possibilities of type IV pili or flagella being involved in the coaggregation responses, we sought to determine whether the strains of P. aeruginosa and S. marcescens possessed these motility-mediating surface structures by assessing the motility patterns of the strains used. All P. aeruginosa strains were motile (swimming) by the hanging drop assay, and P. aeruginosa 6294, 6206, and ATCC 9027 exhibited twitching motility by the agar stab assay (Fig. 2). All S. marcescens strains were motile by the hanging drop assay, and three strains (ATCC 13880, 27, and 35) showed swarming motility (Fig. 2). However, there was no overall relation of these motility types and the ability to coaggregate. 
Figure 2
 
Representative images of the twitching and swarming motility of P. aeruginosa (6294, ATCC 9027) and S. marcescens (ATCC 13880, 27). (A, B) The pili-mediated motility of P. aeruginosa; the arrow in (A) indicates a halo zone of pili motility of P. aeruginosa 6294 (halo zone: A>B). (C) The swimming motility of S. marcescens ATCC 13880; (D) the swarming-type motility of S. marcescens 27 with a dendritic pattern.
Figure 2
 
Representative images of the twitching and swarming motility of P. aeruginosa (6294, ATCC 9027) and S. marcescens (ATCC 13880, 27). (A, B) The pili-mediated motility of P. aeruginosa; the arrow in (A) indicates a halo zone of pili motility of P. aeruginosa 6294 (halo zone: A>B). (C) The swimming motility of S. marcescens ATCC 13880; (D) the swarming-type motility of S. marcescens 27 with a dendritic pattern.
Staphylococcus aureus 31 adhered less (7.6 ± 0.2 log10 CFU; colony forming unit) to the surface of contact lens storage cases compared to P. aeruginosa 6294 (8.4 ± 0.1 log10 CFU) when grown in isolation. There was no evidence of positive cohesion between P. aeruginosa 6294 and S. aureus. When S. aureus 31 and P. aeruginosa 6294 were grown together, S. aureus 31 was inhibited irrespective of which bacterium was added first. When both P. aeruginosa 6294 and S. aureus 31 were grown together in lens cases, their recovery was reduced 2.8 log10 CFU and 5.0 log10 CFU, respectively, compared to their viable count when adhered in isolation (Fig. 3). When these strains were examined for the production of substances that could inhibit each other, there was inhibition of the growth of S. aureus when P. aeruginosa was spotted (inhibitory zone size 6.0 ± 0.3 mm) but not when S. aureus was spotted onto a lawn of P. aeruginosa. Representative images of the inhibition for S. aureus 31 and P. aeruginosa 6294 are shown in Figure 4
Figure 3
 
Adhesion of S. aureus 31 and P. aeruginosa 6294 when grown together compared to when grown separately.
Figure 3
 
Adhesion of S. aureus 31 and P. aeruginosa 6294 when grown together compared to when grown separately.
Figure 4
 
Production of inhibitory substances observed as halo zones by P. aeruginosa 6294 on a lawn of S. aureus 31 (I) or S. aureus 31 on a lawn of P. aeruginosa 6294 (II).
Figure 4
 
Production of inhibitory substances observed as halo zones by P. aeruginosa 6294 on a lawn of S. aureus 31 (I) or S. aureus 31 on a lawn of P. aeruginosa 6294 (II).
Discussion
The primary goal of this investigation was to determine whether bacteria commonly isolated from contact lens cases could coaggregate or cohere. The study demonstrated for the first time that coaggregation occurred between P. aeruginosa strains and strains of S. aureus isolated from the eye. Two Gram-positive bacteria, S. aureus and S. epidermidis, also showed sporadic coaggregation. Serratia marcescens strains did not exhibit any significant coaggregation with P. aeruginosa and exhibited sporadic low-level coaggregation with S. aureus only. However, no coaggregating pairs exhibited the high scores given by the positive control pairing of A. naeslundii and S. sanguinis. The coaggregating pairs did not show cohesion; that is, the adhesion of one type of bacterium did not promote the adhesion of the other type from the coaggregating pair. Indeed, this study showed evidence that when grown together, P. aeruginosa could inhibit the growth of S. aureus
Sato et al.34 demonstrated that coaggregation between actinomycetes and streptococci occurred via lectin-like substances (i.e., substances similar or identical to proteins that bind sugars) on the surface of actinomycetes with carbohydrate(s) on the surface of streptococci. The coaggregation of Acintomyces oris with Streptococcus oralis is mediated by a single protein on the surface of the actinomycetes named coaggregation factor A35 or by proteinaceous type 2 fimbriae on the actinomycetes36 interacting with carbohydrates on the streptococci.36 However, in the current study, pretreating staphylococci with lactose or sucrose inhibited the coaggregation between them and P. aeruginosa. Pretreating P. aeruginosa with pronase also inhibited the coaggregation. This demonstrates that the staphylococci have a substance on their surface, presumably a lectin resistant to digestion by the protease from Streptomyces griseus, that interacts with carbohydrates on the surface of P. aeruginosa containing structures similar to lactose and sucrose, as well as a protein on the surface of P. aeruginosa that mediates coaggregation. When the sugars, fucose or galactose, were incubated with P. aeruginosa they inhibited coaggregation with the staphylococci. This latter finding implicates the two specific lectin proteins of P. aeruginosa LecA and LecB (PA-I and PA-II, respectively)37 presumably binding to the carbohydrates on the surface of the staphylococci. Sucrose is composed of glucose and fructose; lactose is composed of glucose and galactose; galactose and fucose are monosaccharides. LecA has been shown to specifically bind to galactose or glucose, while LecB binds specifically to fucose or mannose.38,39 In addition the type IV pili of P. aeruginosa can bind to the b-N-acetylgalactosamine (1–4)-b-galactose via the pilus subunit PilA. Staphylococcus aureus strains can produce various capsular polysaccharides that can coat their surface, and these capsules can contain N-acetylgalactosaminuronic acid, N-acetyl-D-fucosamine, N-acetyl-D-glucosaminuronic acid, and N-acetylmannosaminuronic acid,40 which may represent the ligands for the P. aeruginosa protein(s). Pseudomonas aeruginosa has a number of surface polysaccharides that may be involved in coaggregation including glycosylated PilA,41 flagella,42 and lipopolysacchairde of its outer membrane,43 which can have D-glucose, D-galactosamine,43 or N-acetyl-D-fucosamine44; these are implicated as being potential coaggregation sites based on the inhibition studies reported herein. Thus, in this instance both coaggregating pairs appear to have carbohydrate(s) and protein(s) involved in the coaggregation. Perhaps this is why treating with any of the sugars or pronase only partly reduced coaggregation. The fact that sugars could inhibit coaggregation may mean that they could be used to reduce multispecies biofilm buildup in contact lens cases. It is unlikely that the simple sugars used in the current investigation would be of use as these can be used as food sources by many different types of bacteria. However, nonmetabolizable sugar analogues may be of use to control coaggregation. 
Due to the possibilities of type IV pili or flagella being involved in the coaggregation responses, we sought to determine whether the strains of P. aeruginosa and S. marcescens possessed these motility-mediating surface structures by assessing the motility patterns of the strains used. Type IV pili mediate a form of motility in P. aeruginosa known as twitching motility.45 Flagella mediate both swimming and swarming motility.46 All strains of P. aeruginosa and S. marcescens showed (flagella-mediated) swimming motility, and S. marcescens strains ATCC 13880, 27, and 35 showed swarming motility. As only two of the S. marcescens strains and only the ocular isolates of P. aeruginosa (i.e., three strains) showed coaggregation, this demonstrates that possession of flagella per se was not associated with coaggregation. Three strains of P. aeruginosa (6294, 6206, ATCC 9027) demonstrated twitching motility indicating the possession of functional type IV pili, but these strains were not necessarily those that showed coaggregation with staphylococci (6294, 6206, Paer1), again indicating that possession of functional type IV pili is not a requirement for coaggregation. However, as the type IV pili can be either glycosylated or nonglycosylated and the two types of flagella in P. aeruginosa are differently glycosylated, further work is required to determine exactly the relationship between motility and coaggregation in these bacteria.47,48 
The concept of coaggregation contributing to the buildup of bacterial communities on surfaces is well established, especially in the dental literature.49 Thus we sought to determine whether those bacteria that produced coaggregation were associated with cohesion. Cohesion can also occur when a pioneer species modifies the substratum to encourage adhesion of successor species. The most common bacteria found in contact lens cases are staphylococci, which may then be the pioneer species. Gram-negative bacteria such as P. aeruginosa or S. marcescens are much more rarely isolated from lens cases2,5 and so can be considered as successor species. The data in the current study demonstrated that although there was evidence of coaggregation between staphylococci and P. aeruginosa, these bacteria did not cohere. In fact, this study showed evidence for the production of substance(s) by P. aeruginosa that inhibited the growth of S. aureus. While the nature of the inhibiting substances was not investigated in the current experiments, several inhibitory substances have been reported, including DesB (acyl-CoA delta-9-desaturase), 2-heptyl-4-hydroxyquinoline N-oxide and siderophores, and staphylolysin (also called LasA protease).5052 It may be of value to conduct future experiments on these proteins to determine whether any can prevent the growth of S. aureus in contact lens cases. The fact that the production of the inhibitory substance by P. aeruginosa did not result in higher adhesion when P. aeruginosa was allowed to adhere after S. aureus may indicate that death of the S. aureus strains does not remove them from the substrata and so does not result in unveiling of any new adhesion sites for P. aeruginosa
In summary, this study demonstrated for the first time that ocular isolates of P. aeruginosa and S. aureus could coaggregate but that this may not be related to the buildup of biofilms in contact lens cases, as there was no evidence that the coaggregation was associated with cohesion between the strains. Furthermore, the study confirmed that P. aeruginosa can inhibit the growth of S. aureus
Acknowledgments
Disclosure: A. Datta, None; F. Stapleton, Alcon (F), Allergan (F), Coopervision (F); M.D.P. Willcox, Alcon (F), Allergan (C, F), Cochlear (F), Coopervision (C, F, R), Johnson and Johnson Vision Care (F, R), Ophtecs (C, F, R, S), P 
References
Devonshire P, Munro FA, Abernethy C, Clark BJ. Microbial contamination of contact lens cases in the west of Scotland. Br J Ophthalmol. 1993; 77: 41–45.
Willcox MDP, Carnt N, Diec J, et al. Contact lens case contamination during daily wear of silicone hydrogels. Optom Vis Sci. 2010; 87: 456–464.
Dart JK. Predisposing factors in microbial keratitis: the significance of contact lens wear. Br J Ophthalmol. 1988; 72: 926–930.
Wu YTY, Willcox MDP, Zhu H, Stapleton F. Contact lens hygiene compliance and lens case contamination: a review. Cont Lens Anterior Eye. 2015; 38: 307–316.
Szczotka-Flynn LB, Pearlman E, Ghannoum M. Microbial contamination of contact lenses, lens care solutions, and their accessories: a literature review. Eye Contact Lens. 2010; 36: 116–129.
McLaughlin-Borlace L, Stapleton F, Matheson M, Dart JK. Bacterial biofilm on contact lenses and lens storage cases in wearers with microbial keratitis. J Appl Microbiol. 1998; 84: 827–838.
Morgan PB, Efron N, Brennan NA, Hill EA, Raynor MK, Tullo AB. Risk factors for the development of corneal infiltrative events associated with contact lens wear. Invest Ophthalmol Vis Sci. 2005; 46: 3136–3143.
Gray TB, Cursons RT, Sherwan JF, Rose PR. Acanthamoeba, bacterial, and fungal contamination of contact lens storage cases. Br J Ophthalmol. 1995; 79: 601–605.
Yung AMS, Boost MV, Cho P, Yap M. The effect of a compliance enhancement strategy (self-review) on the level of lens care compliance and contamination of contact lenses and lens care accessories. Clin Exp Optom. 2007; 90: 190–202.
Yung MS, Boost M, Cho P, Yap M. Microbial contamination of contact lenses and lens care accessories of soft contact lens wearers (university students) in Hong Kong. Ophthalmic Physiol Opt. 2007; 27: 11–21.
Pens CJ, da Costa M, Fadanelli C, Caumo K, Rott M. Acanthamoeba spp. and bacterial contamination in contact lens storage cases and the relationship to user profiles. Parasitol Res. 2008; 103: 1241–1245.
Wu Y, Zhu H, Harmis NY, Iskandar SY, Willcox MDP, Stapleton F. Profile and frequency of microbial contamination of contact lens cases. Optom Vis Sci. 2010; 87: 152–158.
Wiley L, Bridge DR, Wiley LA, Odom JV, Elliott T, Olson JC. Bacterial biofilm diversity in contact lens-related disease: emerging role of Achromobacter, Stenotrophomonas, and Delftia. Invest Ophthalmol Vis Sci. 2012; 53: 3896–3905.
Larkin DF, Kilvington S, Easty DL. Contamination of contact lens storage cases by Acanthamoeba and bacteria. Br J Ophthalmol. 1990; 74: 133–135.
Wu PZJ, Thakur A, Stapleton F, Willcox MDP. Lens and cornea Staphylococcus aureus causes acute inflammatory episodes in the cornea during contact lens wear. Clin Exp Ophthalmol. 2000; 28: 194–196.
Leitch E, Harmis N, Corrigan K, Willcox MDP. Identification and enumeration of Staphylococci from the eye during soft contact lens wear. Optom Vis Sci. 1998; 75: 258–265.
Cheng KH, Leung SL, Hoekman HW, et al. Incidence of contact-lens-associated microbial keratitis and its related morbidity. Lancet. 1999; 354: 181–185.
Mayo MS, Schlitzer RL, Ward MA, Wilson LA, Ahearn DG. Association of Pseudomonas and Serratia corneal ulcers with use of contaminated solutions. J Clin Microbiol. 1987; 25: 1398–1400.
Kolenbrander PE. Oral microbial communities: biofilms, interactions and genetic systems 1. Annu Rev Microbiol. 2000; 54: 413–437.
Rickard AH, Gilbert P, Handley PS. Coaggregation between aquatic bacteria is mediated by specific growth phase dependent lectin-saccharide interactions. J Appl Microbiol. 2000; 66: 431–434.
Kolenbrander PE, Andersen RN. Inhibition of coaggregation between Fusobacterium nucleatum and Prophyromonas (Bacteroides) gingivalis by lactose and related sugars. Infect Immun. 1989; 57: 3204–3209.
Gibbson RJ, Nygaard M. Interbacterial co-aggregation of plaque bacteria. Arch Oral Biol. 1970; 15: 1397–1400.
Cisar JO, Kolenbrander PE, McIntire FC. Specificity of coaggregation reactions between human oral streptococci and strains of Actinomyces viscosus or Actinomyces naeslundii. Infect Immun. 1979; 24: 742–752.
Costerton JW. Bacterial biofilms: a common cause of persistent infections. Science. 1999; 284: 1318–1322.
Sauer K, Sauer K, Camper AK, et al. Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J Bacteriol. 2002; 184: 1140–1154.
Hall-Stoodley L, Costerton JW, Stoodley P. Bacterial biofilms: from the natural environment to infectious diseases. Nat Rev Microbiol. 2004; 2: 95–108.
Pratt LA, Kolter R. Genetic analysis of Escheria coli biofilm formation: roles of flagella, motility, chemotaxis and type I pili. Mol Microbiol. 1998; 30: 286–293.
O'Toole GA, Kolter R. Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol Microbiol. 1998; 30: 295–304.
Malik A, Sakamoto M, Hanazaki S, et al. Coaggregation among nonflocculating bacteria isolated from activated sludge. Appl Environ Microbiol. 2003; 69: 6056–6063.
Willcox MDP, Patrikakis M, Harty DM, Loo CY, Knox KW. Coaggregation of oral lactobacilli with streptococci from the oral cavity. Oral Microbiol Immunol. 1993; 3: 319–321.
Cookson AL, Handley PS, Jacob AE, Watson GK, Allison C. Coaggregation between Prevotella nigrescens and Prevotella inter-media with Actinomyces naeslundii strains. FEMS Microbiol Lett. 1995; 132: 291–296.
Glick B, Garber N. The intracellular localization of Pseudumonas aeruginosa lectins. J Gen Microbiol. 1983; 129: 3085–3090.
Baldan R, Cigana C, Testa F, et al. Adaptation of Pseudomonas aeruginosa in cystic fibrosis airways influences virulence of Staphylococcus aureus in vitro and murine models of co-infection. PLoS One. 2014; 9: e89614.
Sato S, Koga T, Inoue M. A possible mechanism for the cellular coaggregation between Actinomyces viscosus ATCC 19246 and Streptococcus sanguis ATCC 10557. J Gen Microbiol. 1984; 130: 1351–1357.
Reardon-Robinson ME, Wu C, Mishra A, et al. Pilus hijacking by a bacterial coaggregation factor critical for oral biofilm development. Proc Natl Acad Sci U S A. 2014; 111: 3835–3840.
He D, Hao J, Zhang B, et al. Pathogenic spectrum of fungal keratitis and specific identification of Fusarium solani. Invest Ophthalmol Vis Sci. 2011; 52: 2804–2808.
Winzer K, Falconer C, Garber NC, Diggle SP, Camara M, Williams P. The Pseudomonas aeruginosa lectins PA-IL and PA-IIL are controlled by quornum sensing and by RpoS. J Bacteriol. 2000; 182: 6401–6411.
Blanchard B, Imberty A, Varrot A. Secondary sugar binding site identified for LecA lectin from Pseudomonas aeruginosa. Proteins. 2014; 82: 1060–1065.
Sommer R, Exner TE, Titz A. A biophysical study with carbohydrate derivatives explains the molecular basis of monosaccharide selectivity of the Pseudomonas aeruginosa lectin lecB. PLoS One. 2014; 9: 1–22.
Riordan KO, Lee JC. Staphylococcus aureus Capsular Polysaccharides. Clin Microbiol Rev. 2004; 17: 218–234.
Nguyen LC, Taguchi F, Tran QM, et al. Type IV pilin is glycosylated in Pseudomonas syringae pv. tabaci 6605 and is required for surface motility and virulence. Mol Plant Pathol. 2012; 13: 764–774.
Schirm M, Arora SK, Verma A, et al. Structural and genetic characterization of glycosylation of type a flagellin in Pseudomonas aeruginosa. J Bacteriol. 2004; 186: 2523–2531.
King JD, Kocíncová D, Westman EL, Lam JS. Review: lipopolysaccharide biosynthesis in Pseudomonas aeruginosa. Innate Immun. 2009; 15: 261–312.
Castric P, Cassels FJ, Carlson RW. Structural characterization of the Pseudomonas aeruginosa 1244 pilin glycan. J Biol Chem. 2001; 276: 26479–26485.
Burrows LL. Twitching motility: type IV pili in action. Annu Rev Microbiol. 2012; 66: 493–520.
Harshey RM, Partridge JD. Shelter in a swarm. J Mol Biol. 2015; 427: 3683–3694.
Gastric P. PilO, a gene required for glycosylation of Pseudomonas aeruginosa 1244 pilin. Microbiology. 1995; 141: 1247–1254.
Nothaft H, Szymanski CM. Protein glycosylation in bacteria: sweeter than ever. Nat Rev Microbiol. 2010; 8: 765–778.
Jakubovics NS. Intermicrobial interactions as a driver for community composition and stratification of oral biofilms. J Mol Biol. 2015; 427: 3662–3675.
Filkins LM, Graber JA, Olson DG, et al. Coculture of Staphylococcus aureus with Pseudomonas aeruginosa drives S. aureus towards fermentative metabolism and reduced viability in a cystic fibrosis model. J Bacteriol. 2015; 197: 2252–2264.
Barequet IS, Habot-Wilner Z, Mann O, et al. Evaluation of Pseudomonas aeruginosa staphylolysin (LasA protease) in the treatment of methicillin-resistant Staphylococcus aureus endophthalmitis in a rat model. Graefes Arch Clin Exp Ophthalmol. 2009; 247: 913–917.
Kim S, Yoon Y, Choi KH. Pseudomonas aeruginosa DesB promotes Staphylococcus aureus growth inhibition in coculture by controlling the synthesis of HAQs. PLoS One. 2015; 10: 1–16.
Figure 1
 
Coaggregation between S. aureus (SA) and S. epidermidis (SE) strains. Positive coaggregation was considered to be ≥30% and is indicated by the line on the graph. Also shown is the percentage coaggregation of the positive control strains: Actinomyces naeslundii (AN) with Streptococcus sanguinis (SS).
Figure 1
 
Coaggregation between S. aureus (SA) and S. epidermidis (SE) strains. Positive coaggregation was considered to be ≥30% and is indicated by the line on the graph. Also shown is the percentage coaggregation of the positive control strains: Actinomyces naeslundii (AN) with Streptococcus sanguinis (SS).
Figure 2
 
Representative images of the twitching and swarming motility of P. aeruginosa (6294, ATCC 9027) and S. marcescens (ATCC 13880, 27). (A, B) The pili-mediated motility of P. aeruginosa; the arrow in (A) indicates a halo zone of pili motility of P. aeruginosa 6294 (halo zone: A>B). (C) The swimming motility of S. marcescens ATCC 13880; (D) the swarming-type motility of S. marcescens 27 with a dendritic pattern.
Figure 2
 
Representative images of the twitching and swarming motility of P. aeruginosa (6294, ATCC 9027) and S. marcescens (ATCC 13880, 27). (A, B) The pili-mediated motility of P. aeruginosa; the arrow in (A) indicates a halo zone of pili motility of P. aeruginosa 6294 (halo zone: A>B). (C) The swimming motility of S. marcescens ATCC 13880; (D) the swarming-type motility of S. marcescens 27 with a dendritic pattern.
Figure 3
 
Adhesion of S. aureus 31 and P. aeruginosa 6294 when grown together compared to when grown separately.
Figure 3
 
Adhesion of S. aureus 31 and P. aeruginosa 6294 when grown together compared to when grown separately.
Figure 4
 
Production of inhibitory substances observed as halo zones by P. aeruginosa 6294 on a lawn of S. aureus 31 (I) or S. aureus 31 on a lawn of P. aeruginosa 6294 (II).
Figure 4
 
Production of inhibitory substances observed as halo zones by P. aeruginosa 6294 on a lawn of S. aureus 31 (I) or S. aureus 31 on a lawn of P. aeruginosa 6294 (II).
Table 1
 
Bacteria Used in the Study
Table 1
 
Bacteria Used in the Study
Table 2
 
Coaggregation Between Gram-Positive and Gram-Negative Bacteria After 2 or 24 Hours of Incubation
Table 2
 
Coaggregation Between Gram-Positive and Gram-Negative Bacteria After 2 or 24 Hours of Incubation
Table 3
 
Inhibition of Coaggregation Using Lactose, Sucrose, or Pronase
Table 3
 
Inhibition of Coaggregation Using Lactose, Sucrose, or Pronase
Table 4
 
Inhibition of Coaggregation Using D-Galactose (0.05 M) and L-Fucose (0.05 M)
Table 4
 
Inhibition of Coaggregation Using D-Galactose (0.05 M) and L-Fucose (0.05 M)
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