McAudi Microscope: Dehydration and Sample Preparation. Due to multiple centrifugation steps that may cause cell loss, it is best to attach the cells directly to a slide or plastic sheet to ensure they dry along with the surface. This method helps maintain cell integrity during the dehydration process. The optical microscope requires specific slide thickness for proper imaging. To achieve this, a thin film is first applied to the slides or plastic sheets, ensuring the cells adhere effectively. Here are some common methods for preparing the films:
(1) One of the simplest techniques involves using a polyethylene formaldehyde (formvar) film. Cleaned slides are immersed in a 0.5–1% polyethylene drenched formalin solution dissolved in chloroform. Once the solvent evaporates, a thin, yellow film forms on the slide. Gently shaking the slide in the solution can help achieve an even coating, which is suitable for most optical microscopy applications.
(2) Another effective method uses Poly-L-lysine, a positively charged polymer. A 0.1 mL drop of 1 mg/mL Poly-L-lysine solution (prepared in filtered water) is placed on the cleaned slide and spread evenly. The slide is then dried in a 45°C incubator. This technique is particularly useful because the positive charge of Poly-L-lysine attracts negatively charged cell membranes, enhancing adhesion.
(3) For the McAudi microscope, a glycerin-protein solution is prepared by mixing 30–40% plasma with 3–55% glycerin and distilled water. This mixture is spread onto a clean glass slide and allowed to dry. The resulting protein membrane is then immersed in 2% glutaraldehyde to fix the proteins. After washing and drying, the membrane is activated by adding plasma before use. It is left in the plasma for 30 minutes, then rinsed with buffer solution and ready for direct cell application.
For digital microscopes, cells are fixed and suspended in filtered water to create a suitable cell suspension. These cells are then applied to a film-coated slide or plastic sheet and placed in a humid, low-temperature environment (around 4°C) for 30–120 minutes to allow attachment. Afterward, unattached cells are gently washed away using a fine pair of forceps and clean double-distilled water. The slides are then dehydrated by placing them in a vessel containing 50% acetone (or ethanol). Over time, the concentration of the dehydrating agent is gradually increased every 3 minutes. Some labs prefer to replace half the solution with 100% dehydrant each time, repeating the process 5–6 times until the concentration exceeds 0.5%. At this point, the cells are properly dehydrated.
Once the critical point drying is complete, the slide or plastic sheet is attached to the sample mount using conductive adhesive for metal coating. The third method also allows for efficient cell attachment to the protein membrane. Cells fixed with low-concentration glutaraldehyde can be applied to the "activated" protein film. The slide is then immersed in 2% glutaraldehyde for 10 minutes, followed by a wash in filtered double-distilled water. Finally, the sample is dehydrated, dried, and coated with metal as described above.
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