Impact of the springtail's cuticle nanotopography on bioadhesion and biofilm formation in vitro and in the oral cavity

Abstract

Springtails (Collembola) have a nanostructured cuticle. To evaluate and to understand anti-biofouling properties of springtail cuticles' morphology under different conditions, springtails, shed cuticles and cuticle replicates were studied after incubation with protein solutions and bacterial cultures using common in vitro models. In a second step, they were exposed to human oral environment in situ in order to explore potential application in dentistry. In vitro, the cuticular structures were found to resist wetting by albumin solutions for up to 3 h and colonization by Staphylococcus epidermidis was inhibited. When exposed in the oral cavity, initial pellicle formation was of high heterogeneity: parts of the surface were coated by adsorbed proteins, others remained uncoated but exhibited locally attached, 'bridging', proteinaceous membranes spanning across cavities of the cuticle surface; this unique phenomenon was observed for the first time. Also the degree of bacterial colonization varied considerably. In conclusion, the springtail cuticle partially modulates bioadhesion in the oral cavity in a unique and specific manner, but it has no universal effect. Especially after longer exposure, the nanotextured surface of springtails is masked by the pellicle, resulting in subsequent bacterial colonization, and, thus, cannot effectively avoid bioadhesion in the oral cavity comprehensively. Nevertheless, the observed phenomena offer valuable information and new perspectives for the development of antifouling surfaces applicable in the oral cavity.

Keywords: bioadhesion; collembolan; hexapods; pellicle; saliva; springtail.

Figure 1.

Cuticular structure of the springtail Tetrodontophora bielanensis (Collembola, Hexapoda). Note the characteristic nanostructure of these hexapods' dorsum with mushroom like pins and small cavities arranged in a comb-like pattern. Preliminary in vitro experiments with human saliva indicated interesting so far unknown phenomena of bioadhesion (transmission electron microscopy analysis). Pellicle formation (*) under in vitro conditions after 30-min exposure in centrifuged and sterile filtered human saliva. In direct contact to the surface, the pellicle is arranged as an electron dense less than 10 nm thick basal layer (blue arrows), partially covered by an up to 200 nm thick loosely arranged pellicle layer. Interestingly, cavities between pits on the springtail's surface are covered by membrane-like structures (white triangle) due to the in vitro pellicle formation. These observations gave the impulse for further experiments.

Figure 2.

Wetting of springtail cuticle by the BSA solution. (a) SEM image of an exuviae exposed to the BSA solution for 4 h. The upper insets show non-clogged (i) and clogged areas (ii). (b) Fluorescence imaging: wetting of collembolan cuticle after 3 h with 2 mg ml⁻¹ BSA in phosphate-buffered saline. The bright areas in the left image (‘reflex’ means reflection) represent the reflection signal of the liquid–gas boundary above a still non-wetted part of the cuticle, whereas the bright areas in the right image represent the fluorescence signal of adsorbed BSA on the wetted cuticle (electronic supplementary material, movie S1).

Figure 3.

Bacterial assays on springtails and their polymeric replicates. Note that the start conditions of the cell density are 100× smaller for the 24 h (6 × 10⁶ cells ml⁻¹) assays than for the short-term assays of 2 min and 30 min (6 × 10⁸ cells ml⁻¹). The distribution of cells per colony is presented in box–whisker plots with half of the data points within the box and 80% within the whiskers. Pink lines and red square dots mark the median and the mean values, respectively. Representative SEM images are given in electronic supplementary material, fig. S1. NS, nanostructural features.

Figure 4.

Pellicle formation on springtails in situ (in the oral cavity). After 3 min (a,b), sometimes the formation of membrane-like structures was observed, covering the cavities between the pits. Partial association of granular and globular protein aggregates with the springtails' surface occurred on top of the pits and within the cavities between pits. In general, the typical continuous electron dense basal pellicle which is typical for the dental pellicle was not detectable unequivocally. After 120 min (c), more pronounced adsorption of granular and globular structures on the pit's surface and within the cavities was observed. However, this was not a homogeneous process. Note adherence of first bacteria (B).

Figure 5.

Initial bacterial colonization of springtails in the oral cavity. After 8 h of oral exposure (a–c), more bacteria were detectable. Note the inhomogeneity of pellicle formation by adsorption of globularly and granularly shaped protein aggregates forming network-like structures. Bacteria are embedded in the pellicle layer or adhere to the pellicle surface. Sometimes, bacteria seemed to prefer the large-scale cavities of the springtail surface.

Figure 6.

Progressing biofilm formation on springtails exposed to the oral fluids for 24 h. The pattern of bacterial colonization varied considerably (a–f). On some samples, membrane-like bridging of the cavities by adsorbed protein layers was observed (a,c,d), combined with very low level of bacterial colonization. Others yielded filling of the cavities by adsorbed protein aggregates (e) and extensive bacterial biofilm formation (f). Irrespective of the bacterial colonization also the ultrastructure of the underlying pellicle varied considerably. Globular and granular structures of differing electron density were observed, and the pellicle was of very different tenacity as indicated by the varying pattern of attachment to the springtails' surface.