Mouse Models of Epididymitis Induced by Pathogen-Associated Molecular Patterns

Summary

Here, two distinct surgical injection models are described for inducing inflammatory stimuli in the initial segment (interstitial injection) and cauda epididymidis (intravasal injection) of mice. These methods enable studies of the region-specific responses in the different regions of the epididymis.

Abstract

Epididymitis is a highly prevalent disease in the outpatient urologic clinic and a relevant factor in male infertility. Bacterial infections, including sexually transmitted bacteria and common uropathogens, represent the most common etiological factors of epididymitis. Nevertheless, the pathophysiological mechanisms underlying the onset, natural history, and outcomes of bacterial epididymitis remain poorly understood. In this regard, rodent models are valuable tools for investigating the mechanistic responses of the epididymis when challenged with various inflammatory or infectious stimuli. Studies in rats and mice showed that bacterial-derived pathogen-associated molecular patterns (PAMPs), such as lipopolysaccharide (LPS) and lipoteichoic acid (LTA) from Gram-negative and Gram-positive bacteria, respectively, trigger acute inflammation in the epididymis. The severity of these inflammatory responses differs according to the epididymal region affected and is associated with poor sperm parameters. Herein, two distinct surgical approaches are described in detail for inducing epididymitis in mice through the injection of PAMPs in the interstitial compartment of the initial segment or the luminal compartment of the vas deferens toward the cauda epididymidis, thus allowing investigation of the inflammatory responses of the epididymal proximal and distal regions. It is expected that these experimental models of epididymitis will enable future studies aimed at advancing the understanding of the mechanisms governing the epididymal responses to invading pathogens, opening novel ways for therapeutic interventions to mitigate the repercussions of epididymitis on male fertility.

Introduction

The epididymis is a mucosal male reproductive organ composed of a single, highly convoluted tubule that connects the efferent ducts to the vas deferens. The epididymis is essential for sperm maturation (acquisition of sperm motility and fertilizing ability), concentration, transport, storage, and protection until the ejaculation^1,2. Classically, the rodent epididymis has four anatomical regions: the initial segment, caput, corpus, and cauda¹ (Figure 1). Furthermore, each region of the epididymis can be subdivided into additional segments separated by interstitial connective septa, indicating the presence of 10 segments in the mouse epididymis³ (Figure 1).

The immunobiology of the epididymis is an expanding field of investigation due to the association between immune dysregulation and male infertility⁴. Studies in the last two decades have unraveled that the epididymis is populated by different components of the immune system, such as resident immune cell types (e.g., mononuclear phagocytes and lymphocytes) and pathogen-recognition receptors [PRRs, e.g., Toll-like receptors (TLRs) and NOD-like receptors] expressed on several non-immune and immune cell types, which together create dynamic region-specific epididymal microenvironments^5,6,7,8,9. These epididymal immune components play a crucial role in the dual ability of the epididymis to coordinate immune tolerance towards immunogenic spermatozoa while rapidly triggering immune responses to ascending urethral infections and other harmful stimuli^4,10. Factors affecting this intricate immune balance in the epididymis warrant further investigation since they can be detrimental to epididymal function and, thus, may negatively impact male fertility.

In this context, inflammatory diseases of the epididymis, collectively known as epididymitis, are highly prevalent in men of all ages and are important etiological factors in male subfertility and infertility^4,11,12. Epididymitis is the fifth most common male urological diagnosis in men between 18 and 50 years old, with ~600,000 cases per year in the US alone^4,13. Ascending canalicular infections with urogenital pathogens (e.g., Escherichia coli) or sexually transmitted diseases (e.g., Chlamydia trachomatis) are the most common cause of epididymitis^4,14. Most epididymitis patients experience pain in the cauda epididymidis but less frequently in the caput region, underscoring the clinical implications associated with the ability of the epididymis to trigger region-specific responses to inflammatory stimuli¹⁰. Epididymitis can cause profound deterioration in seminal parameters (e.g., sperm concentration and motility), together with pronounced leukocytospermia in the acute phase of the disease^4,12,15. Notably, a substantial cohort of epididymitis patients suffers from persistent oligozoospermia or azoospermia, despite successful pharmacological therapy and symptom remission^16,17. The mechanisms underlying the long-term impact of epididymitis outcomes on sperm quality remain poorly understood.

Human acute epididymitis samples are rare, since epididymal biopsies are contraindicated due to the risk of uncontrolled dissemination of pathogens through the puncture, which can lead to sepsis and organ damage, and sperm leakage to the interstitial compartment due to duct rupture⁴. Thus, animal models of epididymitis represent relevant tools to investigate the disease mechanisms, allowing the evaluation of inflammatory responses and outcomes^4,12. In this regard, rodents (rats and mice) represent the most common model organisms used to study epididymitis in a controlled experimental setting^4,12. Different rodent models of bacterial epididymitis have been established based on the type of pathogen (e.g., uropathogenic E. coli and C. trachomatis), infection routes (e.g., retrograde canalicular injection and interstitial injection), local of infection (e.g., initial segment and cauda epididymidis), and temporal post-injection course (e.g., hours to days)^4,12.

The use of pathogen-associated molecular patterns (PAMPs) represents a valuable tool to study epididymitis, since it allows the investigation of pathophysiological mechanisms of the disease triggered by different molecules associated with distinct pathogens (e.g., bacteria, viruses, and fungi) through the activation of specific PRRs, such as TLRs and signaling pathways^{5,18,19,20,21}. For instance, PAMPs derived from the cell wall of Gram-negative and Gram-positive bacteria, such as lipopolysaccharide (LPS; TLR4 agonist) and lipoteichoic acid (LTA; TLR2/TLR6 agonist), respectively, have been used to elicit epididymitis through multiple administration routes, including intravenous, intraperitoneal, or via direct injection into the epididymal regions¹⁰. Under these conditions, both LPS and LTA have been shown to induce inflammatory responses in the epididymis in rats and mice, associated with modulation of a distinct subset of inflammatory mediators and immune cell recruitment in a region- and temporal-specific manner^19,20. In particular, the region-specific administration of PAMPs in the interstitial compartment of the initial segment or in the luminal compartment of the vas deferens toward the cauda epididymidis of mice recently emerged as relevant models to investigate the proximal and distal response of the epididymis to different inflammatory stimuli, thus providing a framework for the understanding of the different immune environments of the organ and their contributions to disease outcomes^4,18,19.

Here, two different experimental protocols are presented for inducing direct inflammatory stimuli in distinct regions of the mouse epididymis, namely the initial segment and the cauda epididymidis, using interstitial and retrograde intravasal injection routes, respectively. Combined, these different approaches permit a regional delivery of PAMPs and the evaluation of opposite regions of the epididymis during inflammation.

Protocol

All animal experiments were performed in accordance with the Guide for the Care and Use of Laboratory Animals (National Institute of Health, USA) and National Council for the Control of Animal Experimentation (CONCEA, Brazil). Procedural approval was granted by the Ethics Committee for the Use of Experimental Animals (CEUA-IBB/UNESP, Brazil; protocol number: 6272050320; 2269040625). Adult C57BL/6 male mice (90-120 days) were used in these studies. The reagents and the equipment used in this study are listed in the Table of Materials.

1. Interstitial injection in the initial segment of the epididymis

1. Mouse preoperative preparation
   1. Induce general anesthesia with ketamine/xylazine (60/20 mg/kg, i.p.) (following institutionally approved protocols). When anesthetized, place the mouse in a supine position on the surgical field.
      NOTE: Before incision, firmly pinch the toe to ensure the pedal reflex is absent. Inhaled anesthetics (e.g., isoflurane) can be used to replace ketamine/xylazine.
   2. Make the trichotomy of the mouse abdomen using the electric trimmer pen shape (Figure 2A).
   3. Perform abdominal asepsis with sterile cotton wet with chlorhexidine 0.25% (v/v) and alcohol.
2. Injection in the interstitial compartment of the initial segment
   1. Using the scalpel size 15, gently make a ~4 mm vertical incision on the right side of the abdominal wall (Figure 2A).
      NOTE: Be careful with the preputial gland.
   2. With the angled blunt-ended surgical forceps, gently separate the skin. Using the scalpel size 15, gently make a ~4 mm vertical incision on the peritoneum at the site of the first incision in the abdominal wall.
   3. Carefully lift the epididymis and testis for the abdominal space, pushing the right side of the scrotum.
      NOTE: To facilitate the following procedures, use either the headband magnifier or a stereoscopic microscope.
   4. With the curve surgical forceps, carefully pull the epididymal white adipose tissue (eWAT) to localize the initial segment of the epididymis. Using the eWAT, gently expose the initial segment at the incision area (Figure 2A).
   5. Position the Hamilton syringe, preloaded with the injection solution, at an angle approximately 90° parallel to segment #1 of the initial segment. Carefully insert the needle in the interstitial space with the needle bevel oriented down (Figure 2A and Supplementary Figure 1).
      NOTE: Be careful not to puncture the epididymal duct; do not tilt the needle.
   6. Position the curve surgical forceps in the needle insert region and gently pinch the epididymal capsule and the needle. Slowly lower the syringe plunger until all the stimulus solution is expelled (Figure 2A).
      NOTE: Maximum injection volume: 10 µL.
   7. Keeping the curve surgical forceps in position, carefully remove the needle. After ~30 s, remove the surgery curve forceps.
   8. Using the eWAT, gently return the tissue to the abdominal cavity.
      NOTE: Do not pinch the initial segment, only the eWAT.
   9. Localize the peritoneum and make two suture stitches with the needle holder and size 5/0 surgical suture (Supplementary Figure 1).
      NOTE: Check that the incision in the peritoneum is closed; if necessary, make another stitch.
   10. Close the skin incision with two suture stitches with the needle holder and size 5/0 surgical sutures (Figure 2A).
   11. Repeat steps 1.2.1-1.2.10 to make the interstitial injection on the left side.
3. Postoperative care
   1. Observe the mouse postoperatively to ensure appropriate recovery (check animal behavior, shivering, locomotion, and other signs of toxicity). Keep animals warm until they recover from surgery. It is expected that the mouse will move more slowly within the cage after surgery, returning to normal behavior after ~12-24 h.

2. Intravasal injection in the cauda epididymidis

1. Mouse preoperative preparation
   1. Induce general anesthesia with ketamine/xylazine (60/20 mg/kg, i.p.). When anesthetized, place the mouse in a supine position on the surgical field.
      NOTE: Before incision, firmly pinch the toe to ensure the pedal reflex is absent. Inhaled anesthetics (e.g., isoflurane) can be used to replace ketamine/xylazine.
   2. Place the index finger and thumb below the scrotum to check the position of the testes and epididymides. If necessary, carefully massage the sides of the abdomen toward the scrotum to position the testicles and epididymides in the scrotum.
   3. Make the trichotomy of the mouse scrotum using the electric trimmer pen shape (Figure 3A).
   4. Perform scrotum asepsis with sterile cotton wet with chlorhexidine 0.25% (v/v) and alcohol.
2. Retrograde injection in the luminal compartment of the vas deferens toward the cauda epididymidis
   NOTE: To facilitate the following procedures, use either the headband magnifier or a stereoscopic microscope.
   1. With the index finger and thumb below the scrotum, keeping the skin of the scrotum gently stretched, gently make a ~2 mm vertical incision on the medial portion of the scrotum with the scalpel size 15, cutting the skin, tunica dartos, and external spermatic fascia (Figure 3A).
   2. Using the index finger, gently move the left testis and epididymis back to the abdomen, keeping the right testis and epididymis in the scrotum.
   3. With the index finger and thumb below the right testis and epididymis, keeping the skin of the scrotum gently stretched, carefully open and close the angled blunt-ended surgical forceps on the internal spermatic fascia, until breaking the internal spermatic fascia and the vaginal tunic.
   4. Relieve pressure from the index finger and thumb. Through the opening in the vaginal tunic, insert the angled surgical forceps towards the base of the scrotum. Carefully pull the distal epididymal fat to localize the cauda epididymidis.
   5. Localize the vas deferens and position the straight surgical forceps below the vas deferens. In the vas deferens, position the bulldog tweezer clamp, keeping enough space for opening the straight surgical forceps that must be between the cauda epididymidis and the bulldog tweezer clamp (if necessary, limit the opening of the straight surgical forceps to avoid causing damage to the vas deferens) (Figure 3A).
   6. Position the Hamilton syringe, preloaded with the injection solution, at an angle of approximately 10-15° parallel to the vas deferens and insert the needle bevel oriented up (Figure 3A, and Supplementary Figure 1).
   7. Squeeze the straight surgical forceps to loosen the vas deferens, and then slowly lower the syringe plunger until all the stimulus solution is expelled (Figure 3A).
      NOTE: Maximum injection volume: 10 µL.
   8. Remove the straight surgical forceps. Position the curved surgical forceps in the needle insert region, pinch gently, and carefully remove the needle.
   9. After ~30 s, remove the surgery curve forceps and the bulldog tweezer clamp. Return gently the vas deferens to the scrotum.
   10. Localize the vaginal tunic and make one suture stitch with the needle holder and size 5/0 surgical suture (Supplementary Figure 1).
       NOTE: Check that the incision in the vaginal tunic is closed; if necessary, make another stitch.
   11. Using the index finger, gently move the right testis and epididymis back to the abdomen and position the left testis and epididymis in the scrotum.
   12. Repeat the steps 2.2.3-2.2.10 to make the intravasal injection on the left side.
   13. Close the skin incision with two suture stitches with the needle holder and size 5/0 surgical sutures (Figure 3A).
3. Postoperative care
   1. Observe the mouse postoperatively to ensure appropriate recovery (check animal behavior, shivering, locomotion, and other signs of toxicity). Keep animals warm until they recover from surgery. It is expected that the mouse will move more slowly within the cage after surgery, returning to normal behavior after ~12-24 h.

Results

Here, two distinct surgical injection models for inflammatory stimuli in the initial segment (interstitial injection) and cauda epididymidis (intravasal injection) in mice are described. The former triggers inflammatory responses in the proximal epididymis, whereas the latter triggers them in the distal epididymis.

The interstitial injection is performed through an incision in the abdomen, making use of the possibility of moving...

Discussion

Here, two distinct surgical injection procedures for the induction of region-specific epididymitis in mice are presented. They are used to perform experimental models of epididymitis specifically targeting the initial segment via interstitial injection or the cauda epididymidis via retrograde intravasal injection. The detailed and visual guide increases the clarity and accessibility of these protocols, seeking to enhance the reproducibility of these methods.

The diameter of t...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Angled blunt-ended surgical forceps	Bonther	xtycga1012
Angled surgical forceps	Bonther	pwghvv1012
Blue Evans solution	Sigma-Aldrich	E2129
Bulldog tweezer clamp	Bonther	jtkpqs1012
Chlorhexidine	Rioquímica	Riohex 0.25%
Curve surgical forceps	Bonther	506e6e1012
Electric trimmer pen shape	-	-
Hamilton syringe	Hamilton	80400
Headband magnifier	OptiVISOR	-
Ketamine	Venco Animal Health	Ketalex
LPS  from E. coli 055:B55	Invivogen	Tlrl-pb5lps
LTA from Staphylococcus aureus	Invivogen	Tlrl-pslta
Needle holder	Bonther	8pihbf1012
Size 5/0 surgical suture	Shalon Medical	n550cti20
Sterile saline	-	-
Straight surgical forceps	Bonther	ejaiuz1012
Xylazine	Syntec	Xilazin