Visualization of Gut Microbiota-host Interactions via Fluorescence In Situ Hybridization, Lectin Staining, and Imaging

This streamlined protocol details a workflow to detect and image bacteria in complex tissue samples, from fixing the tissue to staining microbes with fluorescent in situ hybridization.

Microbes that are host associated make up this very complex ecosystem that really require imaging to be understood. And through this particle, we're gonna show you how to go through the staining process and achieve visualization of the host microbiome interface. Understanding the biology of host-associated bacteria requires an understanding of their localization within micro environments.

And this technique will enable us to visualize individual taxa within host environments, as well as within the context of other bacteria. Through this technique, it's going to be possible to determine which bacteria may have penetrated through the host tissue, as well as to determine whether key features such as the host mucus may be depleted. Prepare fresh methacarn in a compatible container with 60%absolute methanol, 30%chloroform and 10%glacial acetic acid.

Using sharp and clean tools, cut intestinal segments from the mice for imaging. Minimize disturbing the sample as much as possible and handle the sections by their ridges to avoid affecting the imaging area. Fix the sample as soon as possible after dissection to prevent degradation.

Place the intestinal sections in histology cassettes by delicately holding an edge of the tissue with tweezers. Close the cassette and completely submerge it in fresh methacarn solution, ensuring that the solution is not older than a few hours upon immersion of the cassettes. For clinical samples that will be collected in a clinical suite without access to a fume hood, use a polyethylene storage container with a flap cut into the lid that will permit the passage of the histology cassettes.

Tape this flap shut when not passing samples to prevent the escape of toxic fumes. Place the paraffin in a heat-resistant container and melt in an oven at 60 degrees Celsius overnight. Wash the tissue by pouring out the liquid in the appropriate waste receptacle and incubating it sequentially in absolute methanol, absolute ethanol and xylene as described in the text manuscript.

Open the cassettes slightly to allow the paraffin to enter without losing the tissue segments using double gloves or tweezers. Submerge and close the cassettes in the container of melted paraffin. Place the container back into the 60 degree Celsius oven, ensuring that the cassettes are filled with paraffin and no large air bubbles remain.

After incubation for two hours, remove the container from the oven. Using forceps, carefully remove the cassettes. Heat the oven to 60 degrees Celsius and prewarm a Coplin jar.

Add enough volume of xylene in a glass bottle to cover the glass slides in the jar twice and place parafilm around the lid to prevent evaporation of the xylenes. Allow the temperature of the xylenes to reach 60 degrees Celsius. Prepare the FISH hybridization solution as mentioned in the text manuscript.

Place the slides in the Coplin jar, ensuring that the sections did not come in contact with other slides or the jar and bake the slides at 60 degrees Celsius for 10 minutes. In the fume hood, fill the Coplin jar with prewarmed xylenes from the oven, taking care not to pour directly on top of the samples and potentially dislodge the tissues. Place the Coplin jar back in the 60 degree oven.

Pour used xylenes into a proper waste container, taking care not to disturb the tissue sections on the glass slides and using a pair of forceps to keep the slides from falling out of the Coplin jar. Replenish the Coplin jar with the remaining xylenes and incubate for 10 minutes at room temperature in the fume hood. Incubate the sections in 99.5%ethanol for five minutes at room temperature.

After incubation, remove the slides from the Coplin jar. Wipe the back of the slides on a laboratory wipe or a paper towel and briefly air-dry until the ethanol droplets are gone. Create very close circles around each tissue section using a liquid blocker or PAP pen to limit the area of expansion needed to be covered by the hybridization solution, avoiding ink contact with the section.

Prepare the hybridization solution and add 0.5 micrograms of probe for every 50 microliters of the solution used. Pipette the solution onto the sections on the slide. Overlay the section with flexible plastic coverslips, ensuring that the volume of liquid used covers the entire section.

Create a humid chamber with a pipette tip box with wipes or paper towels that have been soaked with excess hybridization solution or PBS to provide humidity. Incubate the slide in the humid chamber at 45 to 50 degrees Celsius for at least three hours depending on the probe set to reduce evaporation. Remove the plastic coverslips and incubate the slides in FISH washing buffer in a Coplin jar both prewarmed to 50 degrees Celsius.

Place the Coplin jar back into the 50 degree Celsius oven for 10 to 20 minutes. Remove the FISH washing buffer and replace it with PBS in the Coplin jar. Immediately after refilling the Coplin jar with PBS, decant the PBS.

Remove slides from the jar and pipette the counterstain on top of the entire section, while making sure to not touch the tissue with the pipette tip. Incubate at four degrees Celsius for 45 minutes. Wash the stains three times quickly with fresh PBS.

Wipe the back of the slides against a wipe or paper towel and let most of the PBS evaporate off the sections aided by a vacuum line connected to a pipette tip. Mount the sections using a mounting medium. Affix the coverslips to the slide by painting along the edges of the coverslip with clear nail polish, taking care to stay away from the edge of the slide.

Let it set at room temperature. Images of distal colon of mouse mono-colonized with Muribaculum intestinale and the one bi-colonized with Muribaculum intestinale and Bacteroides thetaiotaomicron are shown here. The samples were stained with a FISH probe three prime cyanine-3 tagged specific for Muribaculum isolate and also counterstained with the rhodamine-bound lectin UEA-1 and DAPI.

The images of sections stained with cyanine-3 FISH, DAPI, and combined cyanine-3 FISH and DAPI channels show the localization of Muribaculum intestinale. All bacteria-shaped and bacteria-sized DAPI signals are labeled with cyanine-3 in the mono-colonization state. In the bi-colonization state, in addition to these cyanine-3 and DAPI double positive cells, there are DAPI-stained bacterial cells that are cyanine-3 negative as expected.

With the exception of longer filamentous bacteria, larger DAPI-positive structures are plant material or nuclei from host cells. An intestinal segment that has been sectioned at a depth that provides a view of the lumen as well as longitudinal views of the epithelium and a shallow section segment revealing only cross-sections of crypts and no constant mucus layer or bacteria proves that shallow sectioning will not provide luminal slices of the intestine. Uneven tissue or coverslip coverage results in blurry and unevenly illuminated tile scans.

An example of normal background and high background are also shown here. As we learn more about the microbiota, it's become really obvious that bulk assays such as sequencing are not enough to really determine what happens between different microbial species and how they interact together. And for this, we really require imaging.

This type of technique has allowed some really important discoveries, for example, that deprivation of fiber from the diet causes thinning of the mucus layer and potentially infection by pathogens such as studies from the Martins group, as well as studies into the effects of osmotic diarrhea and how the depletion of the mucus layer really impacts both the host and the microbiota. And these were studies done by the Sonnenberg group, as well as our group.