Visualization of Bacteria in Bladder Biopsy Sections via Fluorescence In Situ Hybridization

Abstract

Source: Neugent, M. L. et al., Detection of Tissue-resident Bacteria in Bladder Biopsies by 16S rRNA Fluorescence In Situ Hybridization. J. Vis. Exp. (2019)

This video demonstrates the visualization of bacteria in deparaffinized bladder tissue from a urinary tract-infected patient. Fluorescently-labeled probe hybridized with bacterial 16s ribosomal RNAs reveals the tissue-associated bacteria and their distribution, providing valuable spatial insights into their density within the sample.

Protocol

All procedures involving sample collection have been performed in accordance with the institute's IRB guidelines.

1. Tissue Preparation for Fixation and Paraffin Embedding

NOTE: Biopsies were taken from consenting women undergoing cystoscopy with electro-fulguration of trigonitis for the advanced management of recurrent urinary tract infection (rUTI). rUTI is defined as 3 UTIs in 12 months. Biopsy collection was performed in the operating room while the patient was under anesthesia after obtaining informed patient consent. All samples were coded and de-identified before experimentation.

1. Obtain a cold-cup (~1 mm³) biopsy from the bladder trigone with urologic forceps via flexible cystoscope and place immediately into a sterile 2 mL cryovial containing 1.5 mL of 4% v/v paraformaldehyde (PFA) prepared in 1x sterile phosphate-buffered saline (PBS).  
   NOTE: 4% PFA in 1x PBS can be made in advance and stored at -20 °C until needed.
2. Fix biopsy for 6 h at room temperature or for 16-24 h at 4 °C.
   NOTE: Fixation duration should be calculated based on the size of the tissue sample and over-fixation should be avoided. It is not advised to use glutaraldehyde as a fixative as it introduces autofluorescence.
3. In a sterilized hood or biosafety cabinet, remove the fixative by pipetting and replace it with 1.5 mL of sterile 1x phosphate-buffered saline (PBS). Keep the samples at 4 °C overnight or immediately perform tissue processing and paraffin embedding.
4. Use a sterilized microtome to section tissue blocks at 5 μm thickness and adhere paraffin tissue sections to charged glass microscope slides. A minimum of two slides per biopsy will be required for the 16S ribosomal RNA (16S rRNA) fluorescence in situ hybridization (FISH) protocol.
   NOTE: The biopsy tissue should be cross-sectionally arranged relative to the cutting plane in order to ensure visualization of urothelial layers in all sections. It should also be noted that section thickness should be optimized for bacterial community detection. Thinner sections or sampling of multiple serial sections may be required in the case of infections where tissue-resident bacteria are extremely scarce (e.g., <1 tissue-resident bacterium per 1,000 mammalian cells).

2. Fluorescence In Situ Hybridization with Universal 16S rRNA Probes

NOTE: Two slides per biopsy are required. One slide is needed for the universal 16S rRNA probe and one slide for a control probe with a scrambled sequence. This is important for distinguishing the true signal from the background signal during microscopy since the bladder epithelium is auto-fluorescent in multiple channels. In addition to the scramble probe, blocking with 0.1% Sudan Black B prior to mounting may reduce background autofluorescence inherent in the tissue.

1. Preparation of reagents and fume hood
   1. Clean an empty fume hood (or appropriately fitted biosafety cabinet) with 70% ethanol.
   2. Prepare the hybridization buffer comprised of 0.9 M sodium chloride (NaCl), 20 mM Tris (hydroxymethyl) aminomethane-hydrochloride (Tris-HCl, pH 7.2), 0.1% sodium dodecyl sulfate (SDS) in 10 mL of sterile-filtered water. 
      NOTE: Sterile-filtered water may be prepared by passing autoclaved distilled water through a 0.22 μM filter. The hybridization buffer can be stored at room temperature, but the SDS may precipitate from the solution. If SDS precipitates, warm the solution in a 50 °C water bath prior to use.
   3. Prepare at least 100 mL each of 95% and 90% ethanol in sterile-filtered water in washed, autoclaved bottles.
   4. Dissolve fluorescently labeled lyophilized probes in 10 mM Tris-HCl (pH 8.0) and 1 mM ethylenediaminetetraacetic acid (EDTA) buffer (TE) prepared in filter-sterilized nuclease-free water to a final concentration of 100 μM. Prepare a dilution of 1 μM in TE buffer for use in this protocol. Store both 100 μM concentrated stock and 1 μM stock reconstituted probes at -20 °C protected from light.    
      NOTE: Do not dissolve fluorescently labeled probes in water. Buffering is required to prevent hydrolysis of the N-hydroxysuccinimide (NHS) ester bond conjugating the fluorophore to the nucleotide probe.
   5. Clean five Coplin jars with 70% ethanol (EtOH), allow to dry, label as follows: xylenes I, xylenes II, 95% EtOH, 90% EtOH, double-distilled water (ddH₂O), and fill with 100 mL of the appropriate solution. This will help avoid confusion in later steps.
2. Tissue de-paraffinization and rehydration
   1. In the hood, place two slides per biopsy into a vertical slide rack.
   2. Place a vertical slide rack into the xylenes I Coplin jar (containing 100 mL of xylenes) for 10 min.
   3. Remove the slide rack from xylenes I and blot the bottom on a paper towel to remove excess xylenes. Place into the xylenes II Coplin jar for 10 min.
      NOTE: Never work with xylenes outside of a certified fume hood.
   4. Rehydrate deparaffinized tissue sections in successive ethanol washes (95% and 90%) for 10 min each in the respectively labeled Coplin jars.
      NOTE: During this stage, warm the Hybridization Buffer to 50-56 °C in a water bath.
   5. Remove the slide rack from the 90% ethanol wash, blot the bottom on paper towels to remove excess ethanol, and place in the ddH₂O Coplin jar containing 100 mL of filter-sterilized ddH₂O for 10 min.
   6. While waiting for the ddH₂O wash, dilute the probes to 10 nM in a hybridization buffer to create the staining solution. Prepare 150 μL of staining solution per slide.  
      NOTE: Protect the staining solution from light by wrapping the tubes in aluminum foil and storing them in a drawer. When working with fluorescently labeled probes on the benchtop, consider turning off overhead lights when able. Probes diluted in the hybridization buffer should not be re-used.
3. Hybridization and counter-staining
   1. Prepare a humidifying chamber for each probe by placing soaked, crumpled Kimwipe and sterile water in the reservoir of a P1000 tip box. Place the tip holder cartridge on top – this is where the slides will sit.
      NOTE: It is important to use a humidifying chamber to prevent the biopsy sections from drying out during hybridization. It should be noted that commercially available humidifier chambers are designed to maintain a stable, humid atmosphere. However, the technique detailed here sufficiently controls humidity at substantially less cost.
   2. Remove the slides from the slide rack and place them onto a fresh paper towel (tissue-side up). Use a Kimwipe to dry the slide. Be careful to only gently dab near (not on) the biopsy section to wick away water. Using a hydrophobic pen, draw a border around the biopsy section and place the slide tissue side up in the humidifying chamber.       
      NOTE: Work quickly so that the tissues do not dry out before hybridization.
   3. Place the humidifying chamber into an incubator set to 50 °C. Pipette 50-150 μL of the staining solution directly on top of the tissue so that the rectangle made by the hydrophobic border around the tissue is filled. Be careful to not add too much solution to overflow the hydrophobic border. Close the box gently.
   4. Incubate overnight (~16 h) at 50 °C in the dark. If the incubator has a window, cover it with aluminum foil to create a dark environment.
      NOTE: A hybridization temperature below the melting temperature of the FISH probe is required for a reliable signal. 50 °C is the optimal temperature for the universal 16S rRNA probe but may not be optimal for other probes.
   5. The following morning, prepare at least 500 mL of Wash buffer comprised of 0.9 M sodium chloride (NaCl), and 20 mM Tris-HCl (pH 7.2) in ddH₂O and filter-sterilize into a sterile bottle with a vacuum bottle-top filter. Warm to 50-56 °C in a water bath.
   6. Remove the slides from the humidifying chambers and carefully wick away any remaining hybridization solution with a Kimwipe. Place the slides in a vertical staining rack.
   7. Place the staining rack into an opaque Coplin jar containing 100 mL of pre-warmed Wash buffer for 10 min. If the Coplin jars are not opaque (e.g., glass), place them in the dark during incubation steps — perhaps under a box or in a drawer.
   8. Repeat the wash step twice with fresh Wash buffer in new Coplin jars.
   9. During the wash steps, prepare the counter-stain by diluting a 100 μg/mL stock solution of Hoechst 33342 1:1,000 in the wash buffer. To the same tube, add Alexa-555 wheat germ agglutinin (WGA) to a final concentration of 5μg/mL and Alexa-555 Phalloidin to a final concentration of 33 nM. Store in the dark until ready for use.         
      NOTE: Hoechst, Alexa-555 WGA, and Alexa-555 Phalloidin label deoxyribonucleic acid (DNA), mucin/uroplakins, and actin, respectively, and may be stored long-term in the dark per the manufacturer's instructions. Fluorescent labels used for probes and counterstains can be customized for the filter sets available for the microscope to be used.
   10. Remove the slides from the last wash and gently wick away excess wash buffer with a Kimwipe. Place the slides tissue-side up on a paper towel and add 50-150 μL of counter-stain directly on top of the tissue so that the hydrophobic border is filled, but not overflowing. Cover up to four slides with a cryo box top and incubate for 10 min at room temperature.
   11. Place the sides back into the staining rack and wash twice more in Coplin jars with fresh Wash buffer, for 10 min each.
   12. Thoroughly dry the slides after the last wash and place tissue-side up on a paper towel under a cryo box top. Squeeze one drop of mounting media directly on top of the tissue. Gently place an appropriately sized coverslip (will depend on biopsy size) on top. Gently press out any bubbles as they will interfere with imaging and allow the cover-slipped slides to cure overnight in the dark.
   13. The next day, seal the edges of the coverslip to the slide with a light coat of clear nail polish. Allow to dry for 10 min in the dark and then store in the dark at 4 °C for confocal microscopy.

3. Visualization of 16S rRNA FISH by Confocal Microscopy

NOTE: For this protocol, best results are achieved with a laser-scanning confocal microscope with 63x and 100x objectives. Proper filter sets for visualization of Hoechst, Alexa-488, and Alexa-555 fluorescence are required. However, standard fluorescent microscopy can be used if a confocal microscope is unavailable. This protocol is for a laser scanning confocal microscope.

1. Switch on the confocal microscope and the computer software associated with the microscope per the manufacturer's instructions.
2. Load the slide and visualize with the 10x objective in the blue (Hoechst) channel. Focus carefully until nuclei are visible.
3. Once focused, change the objective to high magnification (63x or 100x). Add oil on top of the coverslip. Refocus with the new objective, making sure that the objective lens comes in contact with oil while focusing.
   NOTE: Use only fine focusing at high magnification (63x or 100x). Oil should only be used for an oil objective.
4. Quickly assess each slide through the eye-piece in the green (enhanced green fluorescent protein, eGFP/Alexa-488) channel to determine which slides are FISH positive and which are FISH negative.
   NOTE: It is best if this initial assessment/scoring is done blinded by a separate individual to reduce experimental bias.
5. To image the stained biopsies, start with a FISH positive slide and switch to the computer visualization mode. Select the 405 (Hoechst), 488 (Alexa-488), and 555 (Alexa-555) channels. Set the pinhole using the longest wavelength channel, in this case 555. Find the correct focal plane for visualization of labeled bacteria in the 488 channel. Without changing the focal plane, set the gain, laser power, and offset for each channel such that the signal is not saturated, and the background is not over-corrected. Acquire the image in all three channels.       
   NOTE: Use the same settings for the 488 channel to image every slide in an experiment. The laser power may require adjustment in the 405 and 555 channels if the optimal focal plane for bacterial visualization changes between fields.

Materials

Alexa-555 Phalloidin	Invitrogen	A34055	Staining Actin
Alexa-555 Wheat Germ Agglutinin (WGA)	Invitrogen	W32464	Staining Mucin
Bottle top filters	Fisher Scientific	09-741-07	Sterilization
Coplin Jar	Simport	M900-12W	Deparaffinization/washing
Coverslips	Fisher Scientific	12-548-5M	Microscopy
Ethanol	Fisher Scientific	04-355-224	Rehydration
Ethylenediaminetetraacetic Acid, Disodium Salt Dihydrate	Fisher Scientific	S311-500	TE
Frosted Slides	Thermo Fisher Scientific	12-550-343
Hoechst 33342, Trihydrochloride, trihydrate	Invitrogen	H21492	Staining Nucleus
Hydrophobic marker	Vector Laboratories	H-4000	Hydrophobic barrier
Kimwipes	Fisher Scientific	06-666-11
Oil Immersol 518 F	Fisher Scientific	12-624-66A	Microscopy
Paraformaldehyde (16%)	Thermo Scientific	TJ274997	Fixation
ProLong Gold antifade reagent	Invitrogen	P36934	Mounting medium
Sodium Chloride	Fisher Scientific	BP358-10	Hybridization buffer/PBS
Sodium Dodecyl Sulphate	Fisher Scientific	BP166-500	Hybridization buffer
Sodium Phosphate Dibasic Hepahydrate	Fisher Scientific	S373-500	PBS
Sodium Phosphate Monobasic Monohydrate	Fisher Scientific	S369-500	PBS
Syringe	VWR	75486-756	Sterilization
Tris-HCl	Fisher Scientific	BP152-5	TE/Hybridization buffer
Xylene	Fisher Chemical	X3P-1GAL	Deparaffinization
0.22 micron syringe filter	Fisher Scientific	09-754-29	Sterilization