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Direct staining and microscopy methods

Soon after the Maki roll plate method was widely accepted, other techniques were implemented for more rapid diagnosis of medical device-associated infections (Rushford et al., 1993). These techniques were designed to avoid the delay in diagnosis due to the required overnight incubation of culture media. A simple technique for rapid detection of bacteria attached to intravascular catheters is the application of Gram staining as a complementary test right after culturing has been performed. Gram staining is applied to the full length of short catheters (<6 cm) or the distal tip segment of longer catheters that are removed for analysis by light microscopy (Cooper and Hopkins, 1985). To perform Gram staining of the catheter samples, gentian violet, Gram’s iodine and safranin are poured in different disposable containers. The catheter is immersed sequentially in each staining solution for 10 s and washed under water before immersion in the next staining solution. Stained catheters are examined by conventional light microscopy at a magnification of x1000 (Cooper and Hopkins, 1985). One positive aspect of the use of Gram staining as a preliminary screen is that the morphology of stained bacteria is consistent with the organisms identified by culturing. Furthermore, Gram staining allows detection of bacterial colonization of the external and internal surfaces of the catheter. However, this methodology is limited by the optical properties of plastic catheters under the microscope and the fact that bacterial colonization may not be evident after the catheter had been perturbed for culture screening. Another direct staining method that has been utilized is the staining of catheter tips with orange acridine (Zufferey et al., 1988).

The use of scanning and transmission electron microscopy to examine removed catheters has shown that the adherent mode of microbial growth on medical devices has an effect on recoverability using culture methods and on their inherent antimicrobial resistance. For example, the number of bacteria recovered from a biofilm might be an underestimate and biofilms might show increased resistance to specific antibiotics (Marrie and Costerton, 1984). Briefly, for scanning electron microscopy inspection, sections of catheters are immersed in fixative reagents, such as glutaral- dehyde. Then the catheter sample is “metallized,” dehydrated in ethanol for critical point drying before microscopy examination (Marrie and Costerton, 1984; Cohen et al., 1968). For transmission electron microscopy (TEM), using a scalpel, some material from the catheter surface is scraped and fixed (glutaraldehyde is commonly used in cacodylate buffer) and washed. The sample is then postfixed and dehydrated through acetone washes, stained, and reinforced with evaporated carbon before examination by TEM (Marrie and Costerton, 1984; Locci et al., 1981). Electron microscopy allows visualization of biofilms, but it requires additional methods to identify the bacterial species.

Early studies on bacteria contamination of catheters show evidence of significant amounts of glycocalyx, also known as the slime layer or biofilm matrix that covers the cell wall of Gram-positive bacteria. The polysaccharide capsule of Gram-negative bacteria is also termed glycocalyx. In some cases, bacteria-contaminating catheters cannot be cultured or when grown do not form the glycocalyx that is seen in in situ micrographs of catheter surfaces (Marrie and Costerton, 1984; Locci et al., 1981;

Costerton et al., 1981). The extended use of prosthetic devices has a huge impact on coagulase-negative staphylococci as these bacteria emerged as opportunistic pathogens (Christensen et al., 1982). The first report by Bayston and Penny (1972) described the production of glycocalyx by staphylococci and it was then considered a potential indicator of virulence (Bayston and Penny, 1972; Ishak et al., 1985). Therefore, methods for the routine detection of slime production were developed for coagulase-negative staphylococci. The first of these was the Christensen method (Christensen et al., 1982) which involves growing a single colony in a test tube where adherence can be detected by staining of slime. Another widely used method to test for slime production by coagulase-negative staphylococci is the Congo red agar method (Freeman et al., 1989). This method utilizes Congo red, a stain that was previously used to detect the presence of exopolysaccharide of bacilli by optical microscopy.

Imaging methods with probes and dyes, such as fluorescence in situ hybridization (FISH), offer higher resolution to visualize biofilms on explanted devices. However, it requires the use of laser scanning confocal microscopy and optimization of the method for the different sizes and shapes of devices (Xi, 2014).

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