パラフィン包埋と顕微解析の微生物コロニー バイオ フィルムの薄い断面

The overall goal of this procedure is to generate paraffin embedded biofilm sections that can be used to assess the distribution of gene expression as reported by fluorescent protein production within intact native morphologies. This method can help answer key questions in the study of microbial communities such how the expression of specific genes in the production of exopolymeric substances distributed in the context of native biofilm architecture. The main advantage of this technique is that is allows for analysis of gene expression within the preserved morphology of biofilm cross-sections while also being amenable to downstream treatments such as staining of distinct features and high-resolution imaging such confocal or electron microscopy.

After preparing agar tryptone solution according to the text protocol, use a 50-milliliter conical tube to pour 45 milliliters of the solution into a 100-millimeter by 100 millimeter square dish. Allow the agar to solidify for approximately 20 to 30 minutes. Pour a second 15-millimeter layer on top of the first layer, then let it solidify overnight, and if necessary, use a lint-free tissue to remove condensation from the lids.

Next, to spot colony biofilms, after growing an isolated single P.Aeruginosa colony from a frozen stock according to the text protocol, pipette 2.5 to 10 microliters of cell suspension onto a medium bi-layer plate. Colonies may be incubated in the dark at 25 degrees Celsius in 80 to 100%relative humidity for up to four days. Gently pour 15 milliliters of the agar solution over the growth medium and colony and allow the agar to form a gel at room temperature for five minutes.

Then, use a sharp razor blade to cut a square, three-layered chuck with the colony laminated between the top two layers. If preparing multiple samples, ensure that each chuck is cut to a comparable size. Gently remove any excess agar from the colony containing chuck.

Then, wet the flat head of a spatula in PBS or water and gently insert it between the top and bottom layers of agar. Immediately transfer the laminated colony into an embedding cassette labeled with a chemical-resistant marking pen. Place the embedding cassette into a glass slide mailer containing previously prepared fixative.

Process the samples. Use 1X PBS to wash them twice for one hour each. For best results, automate reagent homogenization using the low setting on the spin function of an automatic tissue processor.

To embed the samples, fill a wax mold with molten paraffin wax heated to 55 degrees Celsius, then use a heated flat spatula to quickly transfer the infiltrated chuck from the embedding cassette into the wax mold ensuring that the chuck rests parallel to the base of the mold. Allow the wax to solidify overnight at four degrees Celsius. Samples molded in solid wax may be stored indefinitely at this temperature.

Once the wax has solidified, excise the sample from the mold and use a razor blade to trim excess wax from around the sample. Leave some wax extending from one end of the sample that can be used to clamp it into the microtome. It is important to trim the wax-embedded sample such that its shape facilitates the collection of smooth linear ribbons.

Small imperfections in trimming can produce artifacts in the ribbon and compromise the integrity of the section. Next, heat a water bath to 42 degrees Celsius. Then, clamp the sample into the microtome with the surface of the colony oriented perpendicularly to the edge of the blade.

Trim the sample in 50-micrometer intervals until the desired plane of the colony has been reached from which sections will be collected using a microtome set to a sectioning velocity of 75 to 80 RPM and a clearance angle of six to 10 degrees. Cut a ribbon of the desired number of 10-micrometer-thick sections. Then, use a fine-tipped paint brush to detach the ribbon from the blade.

Now, with forceps or a drop of water at the tip of a Pasteur pipette, gently transfer the ribbon to the water bath. Immediately insert a slide into the water bath and position it below the ribbon at a 45-degree angle. Touch the narrow edge of the ribbon to the slide just below the frosted label.

The ribbon should adhere. Pull the slide out of the water bath and adjust the angle so it's perpendicular to the surface of the water allowing the ribbon to lay flat against the slide along its length. Avoid trapping excess water beneath the ribbon.

Gently stand the slide onto an absorbent lint-free tissue wicking excess water from the section. Then, lay the slide on a paper towel and allow it to dry in the dark at room temperature overnight. Heat a leveled hot plate to 45 degrees Celsius and place the slide on the plate for 30 to 60 minutes.

The wax will become semi-molten and flatten against the slide. Gently lift the slide from the hot plate and lay it flat onto a smooth level room temperature surface for approximately one minute until the wax solidifies. Ensure that the molten wax does not pull to either side of the slide.

With the samples inside a glass slide mailer, de-wax the slides in four washes of clearing agent for five minutes each. Use a Buchner aspirator to remove solution between washes. Use 100%ethanol to wash the slides three times for one minute each.

Then, after rehydrating the slides according to the text protocol, immediately mount the sections in a tris-buffered mounting medium and apply a coverslip. The overlay agar around the section does not react with the fixative, so it does not adhere to the chard slide. This agar can dislodge during rehydration and damage the section, so care should be taken not to agitate the solution during rehydration.

Allow the mounting medium to polymerize at room temperature overnight. Once polymerized, use clear nail polish to seal the coverslip to the slide. Finally, store the sealed slides indefinitely in the dark at four degrees Celsius.

While DIC imaging using a 40 times oil immersion objective can be sufficient to show some morphological features of biofilm thin sections, fluorescence microscopy of strains engineered to constitutively express fluorescent proteins provides enhanced visualization of cellular distribution within the samples. As shown in these panels, images of individual sections can stitched together to generate a cross-section of the entire colony and provides context for the localization of structural features within the overall morphology at the macroscopic level. Using a strain engineered to express fluorescent protein under the control of a specific promoter enables the visualization of the distribution of gene expression.

In addition, colonies can be grown on medium containing dyes, or dyes can be added post-sectioning to stains'specific polysaccharides. Finally, samples can also be prepared and imaged at higher resolution using transmission electron microscopy as presented here. It's important when attempting this procedure to be mindful of the samples'vagility and of the potential to introduce fluorescent and structural artifacts either through fixation, sectioning or rehydration which may influence your interpretation of the data.

This procedure allows for a quantitative image analysis such as correlating the expression profiles of fluorescent reporters with morphological features and the Z-axis of a biofilm. This protocol can be applied to diverse species of biofilm-forming microbes with medical and industrial significance such as Pseudomonas aeruginosa, Vibrio cholerae and Bacillus subtilis. After watching this video, you should have a good understanding of how to grow and prepare colony biofilms for sectioning via paraffin embedding.