Visualization of Candida albicans in the Murine Gastrointestinal Tract Using Fluorescent In Situ Hybridization

The steps described below have been approved by the UCSF Institutional Animal Care and Use Committee (IACUC).

1. Gastrointestinal Colonization of Mice with C. albicans Using Oral Gavage

1. Provide housing for 18−21 g male or female mice (8−10 weeks old) as approved by the local IACUC. Use autoclaved food and water starting on the first day.
   NOTE: The use of sterilized cages, bedding, food, and water reduces the risk of contamination with antibiotic-resistant environmental bacteria that can potentially displace C. albicans from the gut. Further, depending on the experimental question, individual housing of animals should be considered because mice are coprophagic and gut contents will be shared among cohoused animals.
2. The following day, replace the autoclaved drinking water with autoclaved water containing 5% glucose, 1,500 U/mL penicillin G, and 2 mg/mL streptomycin.
3. For convenience, prepare 200x antibiotic stock solutions (Table 1) in advance, filter sterilize, and freeze at -20 °C.
4. Maintain animals on antibiotics for one week.
   1. On days 3 and 6 after antibiotics have been started, assess each animal’s feces for contamination with antibiotic-resistant aerobic bacteria. Holding an animal with one hand, place an uncapped microfuge tube near the anus and collect at least one fecal pellet.
   2. Resuspend each fecal pellet in 1 mL of sterile water and plate 100 µL on Luria broth (LB), yeast extract peptone dextrose (YEPD), or brain heart infusion (BHI) plus blood agar plates. Incubate the plates overnight in a standard incubator (not anaerobic) at 37 °C and assess for bacterial growth.
      NOTE: Plates should exhibit minimal growth (0−20 colonies). If >20 colonies of bacteria are recovered from animals treated with penicillin and streptomycin, then it is unlikely that high-level colonization with C. albicans will be established and the experiment should be aborted.
5. Three days prior to gavage, streak C. albicans from frozen glycerol stocks to YEPD + 2% agar plates and incubate at 30 °C for 2 days.
6. One day before gavage, inoculate one colony of each C. albicans strain to be tested into 5 mL of liquid YEPD. Incubate at 30 °C overnight with shaking.
7. The morning of gavage, dilute the saturated culture of C. albicans to an optical density at 600 nm (OD₆₀₀) of 0.1 in 100 mL of YEPD prewarmed to 30 °C. Shake in an orbital shaker at 200 rpm at 30 °C for between 4 h and 5 h until OD₆₀₀ reaches 1 (mid-log growth).
8. Transfer each 100 mL culture to two 50 mL conical tubes and centrifuge at 3,000 x g for 5 min at room temperature (RT).
   NOTE: C. albicans propagates relatively slowly at RT. If adapting this protocol for another species, the inocula may need to be kept on ice to prevent continued growth.
9. Discard the supernatant, resuspend the pelleted cells in 20 mL of sterile saline per 50 mL of culture, and pool the duplicate resuspended pellets into single conical tubes. Centrifuge at 3,000 x g for 5 min.
10. Discard the supernatant and resuspend in 20 mL of sterile saline. Vortex briefly and remove 20 µL for OD₆₀₀ measurement. Dilute 1:50 in sterile normal saline. Centrifuge the remaining resuspended cells at 3,000 x g for 5 min.
11. Discard the supernatant. For an inoculum size of 1 x 10⁸ CFU, resuspend the pelleted cells to approximately 5 x 10⁸ CFUs/mL in sterile normal saline based on the estimated concentration determined from OD₆₀₀ measurement.
    NOTE: Typically, an OD₆₀₀ of 1 corresponds to ~1 x 10⁷ yeast cells/mL. This value may vary by spectrophotometer and should be verified. If a change in concentration is desired for infections with a different microorganism, plan for an inoculum volume of ≤10 µL/g mouse to minimize discomfort to the animal.
12. Verify the concentration of the inoculum by counting a 1:1,000 dilution with a hemocytometer. Adjust the volume of cells to be gavaged based on the concentration determined using the hemocytometer.
13. Using an animal feeding needle (Figure 1A), gavage each animal with the calculated volume of a given inoculum. See steps 1.13.1−1.13.8 for the gavage procedure.
    NOTE: Training for gavage should be obtained from an experienced practitioner, and the procedure should be mastered in advance. A successful procedure for a single animal typically takes no longer than 1 min.
    1. Attach an autoclaved animal-feeding needle to a 1 mL syringe and fill the syringe with one gavage volume, removing any air bubbles to avoid an inaccurate dose. Place on a sterile surface.
    2. Secure the animal for gavage. Hold the animal at the base of the tail with the dominant hand and slide nondominant hand up the animal’s back to the loose skin just behind the ears.
    3. Using the forefinger and thumb of the nondominant hand, gather the loose skin together tightly; this is the “scruff”. Pick up the animal by its scruff and loop small finger of the same (nondominant) hand around the tail to immobilize animal against the palm of the hand. Gently extend the animal’s head so that its head and body (and therefore neck and esophagus) are in a straight line.
       NOTE: Attempting a gavage while the animal’s body is bent or twisted may result in complications such as esophageal perforation and death of the animal.
    4. Pick up the syringe with the dominant hand and hold perpendicular to the animal, with the curved needle pointing down (Figure 1B). Using the ball of the needle, gently hook an inside corner of the mouth, gently advance the needle along the side of the mouth to the back of the throat, and finally move the needle to the center.
       NOTE: Introducing the needle along a lateral path helps to avoid resistance from the animal’s teeth and tongue.
    5. Once the ball of the needle is at the back of throat, tilt the needle up so that it is parallel with the animal’s body line (Figure 1C).
       NOTE: At this point, the animal will exhibit a gag reflex. This is a good sign that indicates that the needle is positioned correctly, and the gagging motion helps to guide the needle down the esophagus. If there is no gag reflex, the needle may be in the trachea rather than the esophagus. Gently withdraw the needle and proceed back to step 1.13.1.
    6. Allow the animal to swallow the inoculum until only a few millimeters remain visible (Figure 1D). Advance the needle smoothly.
       NOTE: If the needle does not advance, avoid using force, which can damage the esophagus. Sometimes advancing the last half centimeter will require an extra-large gag from the animal. If the needle seems to be stuck, slowly withdraw it and try again from step 1.13.4.
    7. Test that the ball of needle is in the correct location (stomach) by gently advancing the plunger. If there is no resistance, continue delivering the inoculum. Once the inoculum has been administered, slowly withdraw the needle.
    8. Replace the animal in its cage and continue to monitor the animal for 5−10 min for signs of labored breathing or distress. Verify that the animal resumes normal activity soon after gavage.
    9. If the same inoculum will be used with the next animal, clean the needle with an alcohol wipe. Proceed back to step 1.13.1. If a different inoculum will be used, use a new needle and proceed back to step 1.13.1.
       NOTE: It is important to inspect animals the day after gavage. Animals exhibiting signs of distress (e.g., hunched posture, labored breathing) should be humanely euthanized, as they have likely suffered esophageal rupture, damage to the lungs, or another internal injury from which recovery is unlikely.
14. Following gavage of the animals, plate dilutions of the inocula for dose verification.
    1. Prepare 10-fold serial dilutions up to 1:10⁵. Plate one gavage volume of the 1:10⁵ dilution onto a 100 mm plate of Sabouraud dextrose agar.
    2. Incubate the plate for 2 days at 30 °C. Count CFUs to confirm inoculum dose.

2. Preparation of Gastrointestinal Tissues for Histology

NOTE: This section of the protocol is adapted from Johansson and Hansson¹⁶.

1. Prepare 500 mL of methacarn solution (60% methanol, 30% chloroform, 10% glacial acetic acid) in a large screw cap plastic jar for every 2 animals dissected.
   CAUTION: Methanol and chloroform are harmful if inhaled and should be manipulated in a chemical hood. Glacial acetic acid can be corrosive and should be handled with the proper personal protective equipment. Methanol, chloroform, and glacial acetic acid should be discarded as hazardous waste.
2. At the experimental endpoint, euthanize animals humanely according to a protocol approved by the local IACUC.
   NOTE: In antibiotic-treated animals, C. albicans colonization typically ranges from ~10⁶ CFUs/g on day 5 up to ~5 x 10⁷ CFUs/g by day 25.
3. Sterilize dissection tools with 10% bleach and rinse with 70% ethanol.
4. Secure each animal to a dissection surface, with ventral side facing up. Use 30 G needles to secure the limbs to the dissection surface (Figure 2A). Spray the animal with 70% ethanol to clean the fur and minimize sticking to the tools.
5. Using blunt-ended forceps, pinch a section of abdominal skin with nondominant hand. Using scissors with dominant hand, incise the skin and underlying fascia near the base of the pelvis. Extend the incision in a U shape along each side of the peritoneal cavity and up to the rib cage. Lift the skin out of the way.
6. Using a pencil, prelabel histology cassettes for the GI segments to be assessed. For example, for each animal, label separate cassettes as “stomach,” “proximal small intestines,” “mid small intestines,” “distal small intestines,” “cecum,” and “large intestines.” Place a foam pad on the bottom of each cassette (Figure 2A).
   NOTE: Figure 2B outlines suggested sections to place in cassettes.
7. Using blunt-ended forceps, gently extract the cecum from the peritoneal cavity. Use scissors to sever connections between the cecum and the small and large intestines.
   NOTE: Cecal tissue is very delicate and easy to rupture. Take care when manipulating.
8. Place the entire cecum into a histology cassette so that it is untwisted and lays flat (Figure 2C). Place a second foam pad on top and close the cassette. Place the cassette into a screw cap jar containing methacarn.
9. Excise a section of the large intestines that contains 1−2 fecal pellets, with a length less than that of the cassette.
10. Place the tissue section into the cassette, keeping the tissue flat and untwisted. Overlay with a second foam pad and close the cassette. Place the cassette into methacarn.
11. For each section of the small intestines to be sampled, excise a 1−2 cm segment that contains some digestive material.
12. Place the segment into a histology cassette, keeping the tissue flat and untwisted. Overlay with a second foam pad and close the cassette. Place the cassette into methacarn.
13. For the stomach, sever the connections to the esophagus and small intestines. Place in a cassette, keeping the tissue flat and untwisted. Overlay with a second foam pad and close the cassette. Place the cassette into methacarn.
14. Leave the cassettes in methacarn solution at RT for at least 3 h and less than 2 weeks.
    NOTE: There are several points during which the fixed GI segments can be transferred to a commercial histology service. These tissues can be difficult to section without disruption of luminal contents, and optimal results will be obtained by an experienced technician. If the commercial service is willing to perform washes prior to embedding the tissues in paraffin, the cassettes can be sent at this stage along with instructions for step 2.16.
15. Begin melting enough paraffin wax to cover all tissue cassettes in a large beaker in a preheated 70 °C hybridization oven.
16. Following fixation, remove the foam pads and return each intact tissue section back into its cassette. Wash the tissues with the following solutions at RT with shaking: twice for 35 min in 100% methanol, twice for 25 min in 100% ethanol, twice for 20 min in xylene.
    CAUTION: Xylene is flammable, toxic, and can cause damage if inhaled. Use in a chemical hood with proper protection. Dispose of xylene via hazardous waste procedures.
17. Pat the cassettes dry on a paper towel and place in premelted paraffin wax for 2 h at 70 °C in a hybridization oven. Ensure that the cassettes are filled with paraffin and no bubbles remain.
    NOTE: This step allows the wax to infiltrate the GI segment and to stabilize the tissue and contents.
18. Remove the cassettes from wax, allow the excess wax to drain, and store at RT until they are embedded in wax blocks. Discard the wax containing trace amounts of xylenes as hazardous waste.
    NOTE: The Noble Lab uses a commercial histology service for embedding and sectioning of tissues. For standard visualization of C. albicans yeasts and hyphae, 4 µm sections are used. To evaluate connectivity of hyphal segments, which typically extend through multiple planes, 8 µm sections can be used. Depending on the ability of FISH probes to penetrate into the specimen, it may be possible to use even thicker sections.

3. Validation of Newly Designed Probes

NOTE: The following protocol for validating C. albicans probes was adapted from Swidsinski et al.¹⁷.

1. Streak C. albicans SC5314 and closely related comparator species (e.g., Candida dubliniensis, Candida tropicalis) to YEPD + 2% agar plates. Incubate for 1−2 days at 30 °C.
2. Inoculate 5 mL of liquid YEPD medium with a single colony.
3. Pipette 1 mL of the suspended cells to a microcentrifuge tube and centrifuge at 3,000 x g for 5 min. Discard supernatant.
4. Resuspend the pellet in 100 µL of phosphate-buffered saline (PBS).
5. Pipette 5−10 µL of resuspended cells and spot onto a glass slide. Spread into a larger circle and allow to dry.
   NOTE: If cells are not adherent, use poly-L-lysine coated slides.
6. Circle the cell spot with a PAP pen and cover with 50 µL of 2.5% paraformaldehyde (PFA) in PBS. Incubate at RT for 10 min.
   CAUTION: PFA is flammable and can cause health problems if inhaled or in contact with skin. Items contaminated with PFA should be discarded as hazardous waste.
7. Wash 3x with PBS. Tap liquid off onto paper towel and cover with 100 µL of PBS. Repeat twice. Discard final wash.
8. Prepare slides for multiday storage. If performing FISH on the same day, proceed to step 3.9 without the steps in this section.
   NOTE: It is preferable to perform the hybridization immediately but if short on time follow step 3.8 before storing. Slides can be stored for several days at 4 °C.
   1. Cover the spot with 60% ethanol. Incubate at RT for 3 min. Discard the solution.
   2. Cover the spot with 80% ethanol. Incubate at RT for 3 min. Discard the solution.
   3. Cover the spot with 100% ethanol. Incubate at RT for 3 min. Discard the solution. Proceed to step 3.9.
9. Place the slides uncovered in a hybridization oven at 50 °C for 1 h. If step 3.8 was followed, store the slides at 4 °C. If not, proceed to step 4.2 to perform the hybridization protocol.

4. FISH Sstaining in GI Tract Tissues

1. Dewax paraffin-embedded histological sections.
   1. In a hybridization oven, prewarm a Coplin jar filled with xylene to 60 °C.
   2. Insert the slides into the xylene-filled jar. Place at 60 °C for 10 min. Discard the xylene wash. Keep the slides in the jar and pour off the xylene into a waste container.
   3. Pour fresh RT xylene into the jar and incubate at 60 °C for 10 min. Discard the second xylene wash.
   4. Fill the jar with 100% ethanol and incubate at RT for 5 min. Remove the slides from the jar and air dry.
   5. Set the hybridization oven to 50 °C for steps in section 4.2.
2. Stain C. albicans with the FISH probe.
   1. Prepare 1 mL of fresh hybridization solution (20 mM Tris–HCl, pH 7.4, 0.9 M NaCl, 0.1% sodium dodecyl sulfate [SDS], 1% formamide in RNase-free water). See Table 1 for sample calculations.
   2. Mix together 50 µL of hybridization solution with 1 µL of C. albicans probe (5’ Cy3- ACAGCAGAAGCCGTGCC 3’; 50 µg/mL in RNase-free water) for a final probe amount of 0.5 µg per slide.
      NOTE: Many laboratory-bred mice do not contain any fungi among their microbiota. In this case, it is possible to substitute a pan-fungal probe (5’ Cy3-CTCTGGCTTCACCCTATTC 3’) that recognizes 23S rRNA of most fungal species, not just C. albicans. In the authors’ experience, the panfungal probe yields brighter staining than the C. albicans-specific probe.
   3. Protect the solution from light and prewarm to 50 °C.
   4. Using a PAP pen, draw a circle around tissue section from section 4.1.
   5. Pipette 50 μL of probe-containing hybridization solution onto the fixed tissue and gently spread over the tissue section with a pipette tip. Overlay the liquid with a hybridization coverslip making sure to prevent bubbles.
   6. Seal slides in a watertight hybridization chamber and incubate the sections for 3 h at 50 °C in the dark in a hybridization oven.
   7. Meanwhile, prepare 50 mL of FISH washing solution (20 mM Tris-HCl, pH 7.4, 0.9 M NaCl in RNase-free water). See Table 1 for sample calculations.
   8. Move the washing solution to 50 °C to prewarm.
   9. After 3 h, add the washing solution to a Coplin jar. Then, carefully remove the coverslip from the slide with forceps and place the slide in the washing solution.
   10. Cover the jar with aluminum foil and incubate at 50 °C for 20 min.
   11. During this incubation, prepare mucin-nuclei staining solution (20 ng/mL 4′,6-diamidino-2-phenylindole [DAPI], 1.6 µg/mL fluorescein isothiocyanate [FITC]-lectin in PBS). See Table 1 for sample calculations. For stomach and large intestines, use FITC-Ulex europaeus agglutinin 1 (UEA-1). For small intestines and cecum, use a combination of FITC-UEA-1 and FITC-wheat germ agglutinin (WGA).
       NOTE: Lectins from different species recognize different sugar modifications of glycoproteins such as mucin. The lectin combinations recommended here were determined empirically for optimal staining of mucus in different GI compartments.
   12. Wash the slides by pouring off the FISH washing solution and refilling the jar with RT PBS. Pour off the PBS immediately and refill with PBS. Pour off the PBS for a total of 2 washes.
   13. Wick away the PBS with a delicate task wiper tissue and circle the intestinal tissue section with a PAP pen. Place the slides in an opaque plastic container and add 50 µL of mucin-nuclei staining solution to the tissue section. Cover the container with aluminum foil to protect from light. Incubate at 4 °C for 45 min.
   14. Place the slides in a Coplin jar and wash quickly twice in PBS at RT.
   15. Tap away the excess PBS on a paper towel, and then wick away the final drops of PBS with a tissue.
   16. Place one drop of mounting medium on each section and overlay with a glass coverslip, preventing bubbles. Let the mounting medium spread under the entire coverslip. For immediate imaging, anchor the coverslip in place with nail polish at the corners. For long term storage, seal around the coverslip with nail polish.
   17. Image the slides using a fluorescent microscope.