Mouse bladder wall injection is a useful technique to orthotopically study bladder phenomena, including stem cell, smooth muscle, and cancer biology. Before starting injections, the surgical area must be cleaned with soap and water and antiseptic solution. Surgical equipment must be sterilized before use and between each animal. Each mouse is placed under inhaled isoflurane anesthesia (2-5% for induction, 1-3% for maintenance) and its bladder exposed by making a midline abdominal incision with scissors. If the bladder is full, it is partially decompressed by gentle squeezing between two fingers. The cell suspension of interest is intramurally injected into the wall of the bladder dome using a 29 or 30 gauge needle and 1 cc or smaller syringe. The wound is then closed using wound clips and the mouse allowed to recover on a warming pad. Bladder wall injection is a delicate microsurgical technique that can be mastered with practice. Protocol Mouse Bladder Wall InjectionChoice of mouse strain, age, and sex is dictated by experimental needs. We use mice between 8 and 12 weeks of age, since this is a window of immunological maturity prior to senescence. As a general guideline, mice should arrive at least one week prior to experimental manipulation in order to avoid stress-induced confounding factors. Representative Results:A well-localized bleb that does not leak fluid and stays stable in size is an indication of successful injection of the cells into the bladder wall (Figure 1). Histological analysis can be performed to confirm the presence of the injected cells in the bladder wall.

A retrospective cohort study, using the electronic medical records of Kaiser Permanente Northern California (2011)(2012)(2013)(2014)(2015), included 560 robotic and 6785 conventional laparoscopic cases with 1836 "complex" patients (25%). The average operative time was 152 minutes (robotic) vs 157 minutes (conventional) laparoscopic hysterectomy. Complex surgical cases averaged 190 minutes and noncomplex cases averaged 144 minutes. For women with complex disease, the robotic approach, when used by a higher-volume surgeon, may be associated with shorter operative time and slightly less blood loss, but not with lower risk of complications.

Uropathogenic bacterial strains of interest are grown on agar. Generally, uropathogenic E. coli (UPEC) and other strains can be grown overnight on Luria-Bertani (LB) agar at 37°C in ambient air. UPEC strains grow as yellowish-white translucent colonies on LB agar. Following confirmation of appropriate colony morphology, single colonies are then picked to be cultured in broth. LB broth can be used for most uropathogenic bacterial strains. Two serial, overnight LB broth cultures can be employed to enhance expression of type I pili, a well-defined virulence factor for uropathogenic bacteria. Broth cultures are diluted to the desired concentration in phosphate buffered saline (PBS). Eight to 12 week old female mice are placed under isoflurane anesthesia and transurethrally inoculated with bacteria using polyethylene tubing-covered 30 gauge syringes. Typical inocula, which must be empirically determined for each bacterial/mouse strain combination, are 10 6 to 10 8 cfu per mouse in 10 to 50 microliters of PBS. After the desired infection period (one day to several weeks), urine samples and the bladder and both kidneys are harvested. Each organ is minced, placed in PBS, and homogenized in a Blue Bullet homogenizer. Urine and tissue homogenates are serially diluted in PBS and cultured on appropriate agar. The following day, colony forming units are counted. For each mouse bladder and kidney to be sampled, place 5-6 stainless steel beads (bladder, 1.6 mm size or 0.9-2.0 mm blend) or zirconium oxide beads (kidney, 0.5 mm size) in one cryovial tube, respectively. Autoclave the bead-containing tubes. 2. After the tubes cool to ambient temperature, add 200 microliters of sterile PBS to each stainless steel bead-containing tube and 400 microliters of sterile PBS to each zirconium oxide bead-containing tube. 3. We prepare urethral catheters as described by Hung and Hultgren 13 . Autoclave a clean pair of flat-head forceps and fine scissors (iris or similar type) and allow the instruments to cool to ambient temperature. Wipe down the surfaces of a laminar flow hood with 70% ethanol. In the hood, cut a 30 cm (approximate) segment of polyethylene tubing. Aseptically place a sterile 30 gauge needle (1/2 inch long needle) on a sterile 3 cc syringe. Pick up one end of the cut polyethylene tubing with the autoclaved forceps and slide the tubing onto the needle until it meets the hub. Cut the tubing so that approximately 2 cm extends beyond the tip of the needle. Remove the catheter from the syringe and place in a sterile petri dish. 4. To reduce the likelihood of cross-contamination, we make one catheter for each mouse. After all catheters are made, sterilize them by exposure in an uncovered petri dish to UV irradiation for at least 30 minutes. Do not look directly at UV lamps or expose any part of the body in the hood while lamps are activated. After UV exposure, catheters can be stored long-term in the petri dish. We recommend sealing the dish with Parafilm to help maintain sterility.1. Mice should arrive at least a week be...

Objectives This study aimed to estimate the prevalence of pelvic floor disorders by symptoms in female CrossFit athletes in the United States and characterize subjects reporting pelvic organ prolapse symptoms, urinary incontinence, and fecal incontinence. Methods A 27-question anonymous questionnaire was distributed to owners of CrossFit-affiliated gyms. Select questions from validated questionnaires were used to define symptoms. Positive responses with “moderate, or quite a bit” bother defined the presence of urinary incontinence (with stress or urgency). A response of “yes” to “having a bulge or something falling out” defined the presence of pelvic organ prolapse. A response of “yes” to “lose stool beyond your control” questions defined the presence of fecal incontinence. Results Three hundred fourteen respondents had mean age of 36 ± 10 years and a mean body mass index of 25.2 ± 4 kg/m2. Forty-four percent reported ≥1 vaginal delivery. For each workout, respondents reported lifting mean weights of 91 to 217 lb, and 90% reported participation in ≥3 CrossFit workouts per week. Pelvic floor disorder symptoms reported included the following: pelvic organ prolapse, 3.2% (10/314); urinary incontinence, 26.1% (82/314); and fecal incontinence, 6% (19/314). Higher age, parity, and number of vaginal deliveries were associated with urinary incontinence. Higher parity and number of vaginal deliveries were associated with prolapse. Fecal incontinence was not associated with age, body mass index, or obstetric history. Conclusion The prevalence of pelvic floor symptoms in female CrossFit athletes from the general population is likely similar to the general population; however, the prevalence of bothersome urinary incontinence is higher than the general population in women younger than 40 years.