All methods described here regarding animal handling have been approved by the Institutional Animal Care and Use Committee (IACUC) of the Genome Institute of Singapore and Biological Resource Center of the Agency for Science, Technology and Research, Singapore.

1. Mouse Infection

1. Preparation of glass capillaries
   1. Light an open flame source (Bunsen burner or alcohol burner).
   2. Hold a glass capillary with two hands by firmly pinching both ends, then heat the middle of the tube evenly until the glass goes soft. Rotate the capillary gently back and forth along its axis to aid in even heating of the glass.
   3. Remove the glass capillary from the heat source and immediately pull hands apart, while maintaining grip on both ends of the tube. The ideal final length of the pulled capillary is 3-5 cm longer than an unpulled capillary to ensure an appropriate internal diameter for isolating single bladder epithelial cells.
      CAUTION: The tubing still remains extremely hot for a period of time, so set the capillary aside on a heat-safe surface for a few minutes to cool before proceeding to the next step.
   4. Check to see if the middle of the glass capillary has become narrower (Figure 1A) and that the interior of the tube is still hollow (Figure 1B). For isolation of IBCs, a bore size of 200-400 µm is usable.
   5. Pick up one end of the pulled capillary with one hand. Hold a pair of forceps with the other hand, and use it to pick up the pulled glass capillary at its narrowest point. Ensure that the forceps are wielded with enough force to grip the capillary firmly without crushing it.
   6. Use a rapid twisting motion by the hand holding the forceps to snap the pulled capillary at the narrowest point to create a mouth micropipetting capillary.
      NOTE: Using fingers instead of forceps is also acceptable, as long as adequate protection from glass shards is used.
   7. Repeat steps 1.1.2-1.1.6 at least 4x more to produce sufficient spare micropipetting capillaries and provide a range of diameters for IBC isolation. If more than one infection group is anticipated, prepare 5 additional micropipetting capillaries for each additional infection group.
      CAUTION: Do not forget to shut off the open flame.
   8. Place the pulled capillaries in a 100 mm Petri dish and expose the dish to UV radiation for 30 min to sterilize the capillaries.
   9. Replace the lid on the Petri dish after UV sterilization and store the petri dish with the capillaries at room temperature.
2. Preparation of catheters prior to infection
   1. Prepare urinary catheters for infection as described in Hung et al.⁸ and Conover et al.²⁶ at least one day before infection.
3. Preparation of fluorescent uropathogenic E. coli culture
   1. Grow the selected fluorophore-expressing uropathogenic bacterial strain according to established protocols.
      NOTE: The choices of strain and fluorophore largely depend on the microscope and strains available in individual laboratories. In this example, we use a strain derived from UTI89, which is a clinical isolate originally from a patient with recurrent cystitis. This strain, SLC-638, carries a plasmid (pSLC-77) that expresses both vsfGFP-9 and kanamycin resistance¹⁸. SLC-638 is grown in LB broth at 37 °C supplemented with 50 µg/mL kanamycin.
   2. Streak strain SLC-638 on a Luria Bertani (LB)-agar plate supplemented with 50 µg/mL kanamycin. Incubate the plate at 37 °C overnight.
   3. (Optional) View the plate on the dissecting microscope to confirm the expression of fluorescent markers before selecting a colony.
   4. Using a bacterial inoculation loop, transfer the selected colony to a 125 mL conical flask with 10 mL of LB broth supplemented with 50 µg/mL kanamycin. Incubate the flask statically at 37 °C for 24 h.
   5. Subculture the bacteria from this flask by taking 10 µL of culture from the flask and diluting it in 10 mL of LB broth supplemented with 50 µg/mL kanamycin in a fresh 125 mL flask (a 1:1000 dilution). Incubate this second flask statically at 37 °C for another 24 h.
   6. Spin down the bacterial culture for 5 min at 5,000 x g and 4 °C.
   7. Decant the supernatant and resuspend the bacterial pellet in cold PBS at OD₆₀₀ of 0.5.
      NOTE: Although it can vary from culture to culture, typically 1 mL of static culture gives about 4-5 mL of OD₆₀₀ = 0.5 bacterial culture. The total volume of bacterial inoculum needed for each strain can be calculated as follows: 50 µL is required for each mouse and 50 µL is needed for filling the needle head. An additional 10-20% (minimum of 50 µL) of the inoculum is recommended to account for dead volume in the syringe.
   8. Use the remaining bacterial mixture to determine the infection titer as described in Hung et al.⁸.
      NOTE: This step may be delayed for a few hours by storing the bacterial mixture at 4 °C.
4. Murine Model of Urinary Tract Infection
   1. Infect the mice as described by Hung et al.⁸, with one experimental group for each strain of fluorescent E. coli cultured in section 1.3.
      NOTE: Also see Conover et al.²⁶ for visual assistance.
   2. Note the time of bacterial inoculation for the mouse or cage.
   3. Repeat infections for the entire experimental group.
      NOTE: The infection catheter may be reused for all mice in the same group.
   4. Repeat steps 1.4.1-1.4.3 for each experimental group planned, ensuring that a fresh catheter and new lubricant gel is prepared for every group.
      NOTE: For experiments with a large number of animals, it is best to divide the animals into groups of five and staggering the infections such that each group is infected about 30 min to 1 h apart. This will provide enough time for the following steps (Sections 2 and 3).

2. Bladder Epithelial Cell Harvesting to Obtain a Cell Suspension

1. Harvesting and inverting murine bladders
   1. Prepare three 50 mL conical tubes filled with 45 mL of 70% ethanol for sterilizing surgical equipment.
   2. Into two of the tubes prepared in step 2.1.1, place a pair of scissors and a pair of forceps each. Place two pairs of forceps (one preferably narrower and with a rounded tip, for the inversion of the bladder) into the third tube.
      NOTE: The tools in the first tube will be used on the external region, the tools in the second will be used in bladder harvesting, and the two pairs of forceps in the last tube will used in bladder inversion.
   3. At 6 h post infection, euthanize the infected mice according the institution's established IACUC protocols.
      NOTE: Our IACUC protocol calls for euthanasia via cervical dislocation performed while the mouse is under anesthesia (isoflurane).
   4. Lay the animals flat on their backs and use a spray bottle filled with 70% ethanol to sterilize their abdominal area.
   5. Using a pair of forceps and surgical scissors from the first tube (prepared in step 2.1.2), make a small transverse incision on the skin about 1 cm above the urethral opening. Expand the incision diagonally towards the upper limbs of the mouse, creating a V-shaped cut along the entire anterior of the mouse that exposes the contents of the peritoneum. Ensure that during this process, the scissors do not cut through the intestines of the mouse (Figure 2A,B).
   6. Switch to the second set of tools (prepared in step 2.1.2). Using the blades of the scissors or the shafts of the forceps, gently push down on the fat pads near the pelvic region of the mouse.
      NOTE: This step causes the bladder to protrude outward and ensures visibility for harvesting.
   7. Grip the exposed bladder at the apex with a pair of forceps (Figure 2C).
   8. Keeping a firm grip on the bladder apex with the forceps, cut and free the bladder from the rest of the animal (cutting the urethra and ureters away) using the surgical scissors. Do not release the forceps holding the bladder yet.
   9. Switching from the scissors to the narrower rounded forceps from the third conical tube (from step 2.1.2), insert the tip of one shaft of the rounded forceps into the opening of the bladder where it was just cut in the previous step (Figure 2D). With the tip of the rounded forceps safely inserted into the opening of the bladder, release the pair of forceps gripping the apex of the bladder and return it to the second conical tube.
   10. Using the second pair of forceps from the third tube, gently turn the bladder "inside out", first pulling the outside end of the mouth of the bladder away from the rounded forceps (Figure 2D, arrow 1) and guiding it around and over the other tip of the rounded forceps (Figure 2D, arrow 2).
       NOTE: The action can be likened to removing a sock from one foot and pulling it over the other.
       1. During the inversion process, maintain the first rounded pair of forceps almost completely closed. This provides enough freedom of movement to pull the bladder off from the first shaft of the rounded forceps, but also brings the second shaft of the rounded forceps closer to the first and allows for the bladder to be transferred easily. The final result of this step is that the bladder should end up being inverted and on the tip of the second shaft of the first pair of forceps (Figure 2E).
       2. Using the second pair of forceps, gently coax the inverted bladder off the tip of the forceps into 1 mL of cold PBS.
          NOTE: (Optional) This is the opportune time to take images of the entire inverted, infected bladder to observe the general frequency and distribution of IBCs, if any.
   11. Repeat steps 2.1.1-2.1.10 for each bladder in the experimental group (to a maximum of five animals).
2. Bladder epithelial cell scraping
   1. Using two clean pairs of forceps, gently scrape the outside of the inverted bladder (which is the internal epithelial cell layer). The surrounding PBS should appear cloudier as scraping proceeds and epithelial cells are released into the PBS solution.
   2. (Optional) Visually confirm that the bladder scraping has released cells into solution using a dissecting microscope. Figure 3A,B). The PBS should appear cloudy to the naked eye, and the scraped individual bladder epithelial cells can be seen at 10x magnification.
   3. Repeat steps 2.2.1-2.2.2 for each harvested bladder from step 2.1.

3. Intracellular Bacterial Community (IBC) isolation: Mouth Pipetting of IBCs

NOTE: All methods described in this section have undergone an institutional risk assessment. Mouth pipetting carries the inherent risk of ingestion of the solution that is being transferred. This risk is largely mitigated by the nanoliter volumes that this protocol uses, and we recommend that all users of the protocol pay heed to the precautionary and practice notes listed here and in the discussion.

1. After the cells have been scraped into the PBS, set up the mouth micropipetting apparatus (Figure 3C).
   1. Insert the thicker end of the pulled glass capillary (the unpulled end) into the rubber plug (white end) of the aspirator tube.
   2. Insert the narrow end of a 1 mL pipette tip into the other open (red) end of the aspirator tube, ensuring that there is a tight fit.
   3. Insert the narrow end of a 2 mL aspirating pipette into the open, wider end of the 1 mL pipette tip, again ensuring that there is a tight fit.
      NOTE: The resulting setup allows the researcher to mouth pipette from the wider, open end of the aspirating pipette to create a gentle suction force from the narrow end of the capillary tube at the other end of the apparatus.
   4. Test the final mouth micropipetting apparatus using a 100 mm Petri dish containing fresh deionized water. A slight suction action on the open end of the aspirating pipette (similar to sipping on a drink through a straw) should increase the level of liquid in the capillary, but not cause the deionized water to overflow into the aspirator tube. Use one hand to control the capillary tube, while using the other hand to adjust the position of the Petri dish.
      NOTE: The strength of suction required for mouth-pipetting a single IBC will vary among researchers. However, it is recommended that each researcher attempting this technique start with a weak suction and slowly increase it if no liquid is flowing up the capillary. There is no need for a force greater than sucking on a straw for drinking. If the capillary does not appear to be picking up liquid during the test in step 3.1.4, it is possible that either the capillary or the aspirating tube is occluded and needs to be replaced. It is further recommended that all new researchers first practice controlling suction in the mouth pipetting apparatus using sterilized water. Additionally, note that the control of the volume taken up by the mouth pipetting apparatus is through the use of the researcher's tongue. The tongue can finely adjust the strength of suction applied, as well as act as an emergency stop.
   5. After achieving successful uptake of liquid, test the ability to expel it from the capillary by gently blowing into the open end of the aspirating pipette. Ensure that no bubbles are created in the process of expelling the liquid to prevent contamination of the IBCs during step 3.5.
      NOTE: As with suction, the strength of positive pressure applied by the researcher to expel the IBC into the centrifuge tube will vary among researchers. It is recommended for researchers new to this protocol to practice step 3.1 a few days before the actual infection. One suggestion for practicing mouth micropipetting is to practice transferring small volumes of sterilized water mixed with a few drops of food dye (for visibility) using the mouth micropipette apparatus.
2. Place the scraped cell suspension under the dissecting microscope and identify the IBCs as large fluorescent aggregates (Figure 3A,B). The ideal range of magnification is 20-40x. Dip the fine end of the glass capillary into a fresh tube of PBS for 1 s to reduce the uptake of unwanted volume through capillary action.
3. Looking through the microscope, identify the IBC of interest and slowly bring the open end of the capillary tube toward the IBC. Use the fine end of the glass capillary to sweep away extra cells near each IBC to prevent the aspiration of two or more cells, or to break apart larger aggregates of cells.
4. While looking through the microscope, apply a very small suction force on the far end (aspirating pipette) of the mouth micropipetting apparatus to guide the IBC into the glass capillary.
5. After picking up the IBC, move the capillary to an empty 1.5 mL centrifuge tube and apply a slight positive pressure to expel the droplet and the IBC into the centrifuge tube (Figure 3D).
6. Repeat steps 3.3-3.6 on as many IBCs are needed from the current bladder, before proceeding to the next bladder. Change aspirating pipettes and capillaries frequently to prevent build-up of saliva.
   CAUTION: When working with infectious (or clinical) strains of bacteria, constantly monitor the level of solution in the capillary. Do not let the level of liquid being pipetted overflow from the edge of the capillary into the aspirator tube. If this occurs, immediately switch to a different aspirator tube, and discard the previous set up.
7. Repeat steps 3.2-3.6 on all harvested bladders, or until sufficient number of biological samples have been collected for the experimental group.
8. (Optional) Repeat steps 3.6 and 3.7 until all experimental groups (or mice) have been euthanized and sufficient biological samples have been harvested from each group.