Establishment of a Sprague-Dawley rat model of hepatolithiasis based on common bile duct catheterization

Abstract

BACKGROUND

Hepatolithiasis is a common benign bile duct disease prevalent in East Asian regions, characterized primarily by chronic inflammation of the intrahepatic bile ducts, bile stasis, and the formation of calcium bilirubinate stones. The underlying mechanisms of its development have not yet been fully elucidated. Establishing animal models that can simulate the process of chronic cholangitis and the formation of calcium bilirubinate stones is crucial for elucidating the mechanisms of disease development and evaluating potential interventions.

AIM

To establish and optimize a Sprague-Dawley rat model of hepatolithiasis based on common bile duct catheterization combined with staged lipopolysaccharide injection; evaluate its stability, reproducibility, and histopathological characteristics; and provide a reliable animal model for studying the pathogenesis of hepatolithiasis and evaluating related intervention strategies.

METHODS

Twelve specific pathogen-free-grade male Sprague-Dawley rats weighing 400 ± 25 g were randomly divided into an experimental group and a control group, with 6 rats in each group. After establishing a common bile duct catheterization model, the experimental group received lipopolysaccharide (1 mg/kg) injections into the bile duct via the catheter on postoperative days 4, 9, 14, and 19, while the control group received an equivalent volume of saline at the same time points. The catheter was ligated for 6 hours after each injection. On postoperative day 24, the animals were euthanized, and bile and liver tissue were collected. The composition of bile stones was analyzed through gross observation of bile, microscopic examination of bile smears, and Fourier transform infrared spectroscopy. The degree of cholangitis and fibrosis was evaluated using hematoxylin and eosin staining and Scheuer scoring to comprehensively validate the model.

RESULTS

The bile of the experimental group was darker in color and had greater viscosity. Microscopic examination of bile smears showed crystals that resembled human biliary stones. Fourier transform infrared spectroscopy analysis showed calcium bilirubinate to be the major constituent. Hematoxylin and eosin staining showed significant inflammatory infiltration of the portal tracts, and inflammation scores were higher (G1-G2) than those in the control group (G0-G1). The portal tracts in the experimental group showed fibrotic changes (S1), whereas those in the control group did not (S0).

CONCLUSION

This model recapitulates biliary changes, crystal formation, and the process of cholangitis-fibrosis in multiple dimensions. It offers the practicality of long-term drug delivery and periodic sampling through an indwelling catheter, providing a stable experimental model for examining abnormal bilirubin metabolism, biliary inflammatory responses, and the pathogenesis of stone formation.

Key Words: Hepatolithiasis; Sprague-Dawley rat; Lipopolysaccharide; Common bile duct catheterization; Fourier transform infrared spectroscopy; Lithiasis-associated cholangitis and fibrosis

Core Tip: This study successfully established a hepatolithiasis model in Sprague-Dawley rats based on bile duct catheterization and intrabiliary administration of lipopolysaccharide (1 mg/kg) on postoperative days 4, 9, 14, and 19. The model was validated using multiple methods, including microscopic examination of bile smears, identification of calcium bilirubinate crystals by Fourier transform infrared spectroscopy, and histopathological assessment using Scheuer scoring. These findings confirmed that the model consistently produces calcium bilirubinate stones and reproduces typical pathological changes such as chronic biliary inflammation and biliary fibrosis. Additionally, this model allows for repeated intrabiliary dosing and continuous bile sampling, making it a reliable experimental platform for studying the pathogenesis of intrahepatic bile duct stones and evaluating novel intervention strategies.

INTRODUCTION

Hepatolithiasis refers to calculi located at or above the confluence of the left and right hepatic ducts, composed predominantly of pigment-type stones. The disease shows a marked regional distribution, with incidence rates of 2%-30% in China (including Taiwan and Hong Kong), Korea, Japan, Indonesia, and the Philippines, compared with generally below 2% in Western countries[1,2]. Although hepatolithiasis is a benign biliary disease, it has an insidious onset, a prolonged clinical course, and high rates of residual and recurrent stones. It can cause recurrent cholangitis, biliary cirrhosis, and hepatic abscesses and is a strong risk factor for intrahepatic cholangiocarcinoma[1-3].

The prevailing view is that biliary inflammation induced by lipopolysaccharide (LPS), the principal component of bacterial endotoxin, is one of the main causes of hepatolithiasis formation[4,5]. LPS causes inflammatory changes in the biliary epithelium and upregulates the expression of β-glucuronidase (β-G). β-G hydrolyzes conjugated bilirubin (bilirubin diglucuronide) into unconjugated bilirubin and glucuronic acid; unconjugated bilirubin then combines with calcium ions to precipitate as calcium bilirubinate. Biliary epithelial inflammation additionally stimulates the expression and secretion of mucins such as mucin 5AC[6], which increases bile viscosity and further promotes stone formation[5,7,8].

Several animal models have been used to study the pathogenesis of hepatolithiasis, including dietary manipulation alone, single LPS injection combined with common bile duct ligation, short-term repeated LPS injection, and common bile duct catheterization. However, these models generally suffer from short modeling periods, excessively acute inflammation, an inability to allow repeated sampling, and insufficient reproducibility[9-12]. Diet-induced or short-term LPS stimulation models cannot replicate the clinically observed pattern of recurrent cholangitis; complete obstructive models such as bile duct ligation do not reflect the chronic progressive nature of the disease. Moreover, the literature lacks standard operating procedures and systematic model validation systems, inter-experiment variability is large, and it is difficult to determine a consistent chain of causation among biliary compositional changes, stone development, and related histological damage.

Considering these constraints, this study sought to develop a Sprague-Dawley (SD) rat model of hepatolithiasis that recapitulates the clinical disease process while allowing long-term experimentation. SD rats have a stable genetic background, are sufficiently large for surgical manipulation, and have physiological and metabolic properties that resemble those of humans[13]. Common bile duct catheterization created a permanent biliary drug-delivery channel, and the continuous progression of bile stasis, inflammation, fibrosis, and stone-like material was induced through staged intraductal injections of LPS. This work reports a fully documented modeling protocol and systematically tests model reliability using multiple dimensions, including bile composition, crystal structure, Fourier transform infrared spectroscopy (FT-IR) analysis, and histology, thereby providing an experimental basis for further research into the pathogenesis and possible therapeutic targets of hepatolithiasis.

MATERIALS AND METHODS

Table 1 shows the reagents used to establish the SD rat model. Table 2 shows the instruments used to establish the SD rat model.

Table 1 Reagents used during establishment of the Sprague-Dawley rat model.

Reagent	Manufacturer
Lipopolysaccharide	Sigma-Aldrich, United States
Penicillin potassium for injection	Jiangxi Keda Animal Pharmaceutical Co., Ltd.
Isoflurane	Shenzhen RWD Life Science Co., Ltd.
Phosphate-buffered saline	Wuhan Servicebio Technology Co., Ltd.
4% tissue/cell fixative	Beijing Solarbio Science & Technology Co., Ltd.
Environmentally friendly dewaxing/clearing solution	Baisha Bio
Absolute ethanol	Guangdong Guanghua Sci-Tech Co., Ltd.
Liquid Blocker Super PAP Pen (immunohistochemistry pen)	Shanghai Beyotime Biotechnology Co., Ltd.
Hematoxylin staining solution	Baisha Bio
Neutral balsam (long-lasting mounting medium for microscopy)	Shanghai Beyotime Biotechnology Co., Ltd.
Silicone tubing	Jiangsu Juzhuo New Materials Co., Ltd.
Metal guidewire	Asahi Intecc Co., Ltd., Japan
0.9% saline	Kunming Nanjing Pharmaceutical Co., Ltd.
5% glucose saline	Kunming Nanjing Pharmaceutical Co., Ltd.
5-0 Mersilk (Musi) suture	Johnson & Johnson (China) Medical Devices Co., Ltd.
Triangular needle	Hangzhou Aipu Medical Devices Co., Ltd.
Microsurgical instruments	Shanghai Medical Instruments (Group) Co., Ltd.
100-µL microsyringe	Shanghai Gaoge Trading Co., Ltd.
Disposable sterile syringes	Shandong Weigao Group Medical Polymer Products Co., Ltd.

Table 2 Instruments used during establishment of the Sprague-Dawley rat model.

Instruments	Manufacturer
Paraffin microtome	Thermo Fisher Scientific, United States
Water bath	Shanghai Yiheng
Inverted microscope	Thermo Fisher Scientific, United States
KW-10 small/Large animal ventilator	Nanjing Calvin Biotechnology Co., Ltd.
Floor-standing high-speed refrigerated centrifuge	Himac
Multipurpose refrigerated centrifuge	Beckman X-15R
Ultrapure water system	Heal Force Bio-meditech Holdings Limited, China
Ice maker	Sanyo, Japan
Drying oven	Shanghai Yiheng
-80 °C freezer	Thermo Fisher Scientific, United States
-20 °C freezer	Haier, China
Fourier transform infrared spectrometer, NICOLET iS20	Thermo Fisher Scientific, United States

All experimental procedures strictly complied with ethical requirements for animal experimentation and were approved by the Ethics Review Committee of the First Affiliated Hospital of Kunming Medical University (Approval No. 2025DF039). Healthy specific pathogen-free male SD rats weighing 400 ± 25 g were obtained from Henan Skebis Biotechnology Co., Ltd. [Animal License No. SCXK (Yu) 2020-0005]. The animals were housed under specific pathogen-free conditions (temperature, 22 ± 2 °C; relative humidity, 60% ± 10%; 12-hour light-dark cycle) and allowed to acclimatize for 7 days before any procedures. All subsequent steps were carried out strictly according to the experimental plan (Figure 1).

Figure 1 Experimental protocol flowchart. SD male rats were randomized to the control group (n = 6, 1 rat/cage) or the LPS test group (n = 6, 1 rat/cage). Surgery was performed on day 0. LPS or equal-volume saline was injected intraductally on days 4, 9, 14, and 19. All animals were sampled on day 24. Bile was collected for FT-IR analysis and bile smear examination; liver and bile duct tissues were collected for HE-stained paraffin sections. FT-IR: Fourier transform infrared spectroscopy; HE: Hematoxylin and eosin; LPS: Lipopolysaccharide; SD: Sprague-Dawley.

Preoperative procedures

The animals were fasted for 12 hours and deprived of water for 6 hours before surgery.

Surgical instruments: The surgical instruments comprised microsurgical forceps, small curved hemostatic clamps, surgical scissors, microsurgical scissors, sutures and triangular needles (5-0 Mersilk), and a needle holder. All instruments were sterilized by autoclaving at 102.9 kPa and 121 °C.

Additional operative supplies: Additional supplies included disposable sterile drapes, latex gloves, cotton swabs, cotton balls, a dissection board, rubber bands, a heating blanket, a hair clipper, 0.9% normal saline, and 1-mL syringes. All consumables were prepared individually for each rat.

Required materials for catheter preparation: Materials for catheter preparation comprised several 20-cm silicone tubes (inner diameter, 0.3 mm; outer diameter, 0.8 mm), several 4-mm silicone tubes (inner diameter, 0.8 mm; outer diameter, 1.9 mm), a metal guidewire, and a tube dilator. The tip of each 20-cm silicone tube was trimmed to a 45° angle. After dilation, a 4-mm silicone sleeve was fitted around the 20-cm tube approximately 8 mm below its tip, and a 5-0 Mersilk suture approximately 6 cm in length was threaded through the sleeve. A longitudinal side hole was made within 3 mm of the tube tip using a 26-G needle attached to a 1-mL syringe; the hole was made just large enough for the guidewire to pass through. All assembled catheters were sterilized before surgery (Figure 2).

Figure 2 Catheter assembly. A: Schematic of the 20-cm silicone tube tip: 45° bevel cut, side hole within 3 mm of the tip, and 4-mm sleeve mounted 8 mm from the tip with threaded 5-0 Mersilk suture; B: Components from left to right: Finished silicone tube assembly with guidewire, 1-mL syringe with needle, microsurgical curved forceps, and microsurgical needle holder; C: Method for fitting the 4-mm silicone sleeve onto the 20-cm tube; D: Appearance of the completed sleeve assembly; E: Complete catheter tip before insertion, with guidewire in place.

Facility and environmental preparation: The specific pathogen-free operating room was equipped with a sterile operating table, ultraviolet irradiation equipment, and a water bath. The room was disinfected with ultraviolet light for 1 hour before use. Normal saline was warmed to 38 °C. The operating table was covered with a sterile drape, and sterilized instruments were arranged on it. Five 5-cm lengths of 5-0 Mersilk suture were prepared for each rat for intraperitoneal catheter fixation.

Common bile duct catheterization

The anesthesia machine was connected and set to deliver oxygen at 0.8-1.5 L/minute and 4%-5% isoflurane for induction. Induction was completed after approximately 3 minutes, when the righting and toe-pinch reflexes had disappeared. The rat was quickly moved to the dissection board, and anesthesia was maintained via a face mask with oxygen at 0.5-1.0 L/minute and isoflurane at 1.5%-3.0%. The 4 limbs were fixed using rubber bands.

Abdominal hair was removed with a clipper, and the surgical field was disinfected with iodophor. Two small curved vascular clamps were used to lift the skin, and a transverse incision of approximately 2 cm was made about 1 cm below the xiphoid process, slightly to the right. The abdominal wall was opened layer by layer while maintaining muscle tension to avoid injury to the liver and intestines.

Cotton swabs were used to explore the abdominal cavity. The liver was pushed upward and held with a small cotton ball; the intestines were pushed downward and similarly held. The duodenum was exposed, and the white common bile duct, running alongside the portal vein and hepatic artery, was identified (Figure 3A).

Figure 3 Common bile duct catheterization: Operative steps. A: Exposure of the common bile duct (white arrow); the duct runs alongside the portal vein and hepatic artery; B: Blunt dissection of the common bile duct using microsurgical forceps; C: Catheter in situ with initial fixation sutures in place and bile flowing freely from the tube; D: Abdominal closure; the catheter passes through the peritoneum and muscle layer; E: Catheter fixed to the abdominal wall before skin closure; F: External catheter externalized and fixed to the back of the neck, leaving 3 cm of the tube exposed; G: Schematic of a 1-mm-long longitudinal incision in the wall of the common bile duct made using a 26-G needle; H: Schematic of insertion of the silicone tube through the incision in the bile duct using a curved clamp while the guidewire is removed; I: Schematic of the final fixation configuration: Bile duct sutures, sleeve suture to the mesentery, and back fixation suture to the mesentery.

While maintaining appropriate tension, microsurgical curved forceps and cotton swabs were used to bluntly dissect the common bile duct. To avoid injuring the pancreatic duct, dissection was performed at the middle-to-upper segment of the common bile duct (Figure 3B).

Microsurgical forceps were used to gently clamp the lower end of the dissected common bile duct to distend it with bile. A 26-G needle was used to make a longitudinal incision of approximately 1 mm in the bile duct wall, and the silicone tube containing the internal guidewire was introduced through this incision using a small curved clamp, after which the guidewire was withdrawn. Once bile was seen flowing freely from the tube, 2 3-cm lengths of 5-0 Mersilk were used for initial fixation, securing the catheter in situ (Figure 3C). After each fixation step, bile flow was checked to ensure that the sutures were not too tight.

The abdominal cotton balls were removed, the liver and intestines were repositioned, and the cavity was irrigated with 38 °C normal saline and dried with clean cotton balls. A small vascular clamp was used to lift the lower abdominal wall muscle, and microsurgical forceps were passed bluntly from the inside outward through the muscle. The catheter was brought through the peritoneum and muscle layer, confirmed to be tension-free, and fixed to the abdominal wall with 5-0 Mersilk (Figure 3D).

The muscle layer was then closed with continuous 5-0 Mersilk suture, and the catheter was confirmed to be well positioned at the abdominal wall before skin closure (Figure 3E).

The limb rubber bands were released. Hair at the back of the neck was clipped, and the skin was disinfected with iodophor. A small incision was made approximately 0.5 cm behind the interauricular line, just large enough for the catheter. The subcutaneous tissue was bluntly dissected, and the catheter was passed subcutaneously and externalized at the back of the neck, leaving approximately 3-4 cm of the catheter externalized. It was then secured with 5-0 Mersilk suture wrapped 3 times (Figure 3F).

The abdominal skin was closed with continuous 5-0 Mersilk suture and disinfected with iodophor. Six milliliters of 5% dextrose-saline was injected subcutaneously into the back. The rat was transferred to a warming cage and moved to a standard individual cage once fully awake. Each rat was housed individually to prevent tube biting.

For clarity, 3 key technical steps are additionally illustrated schematically: The longitudinal incision made in the bile duct wall using the syringe needle (Figure 3G); insertion of the silicone tube through this incision as the guidewire is withdrawn (Figure 3H); and the final fixation configuration, comprising the bile duct sutures, sleeve suture to the mesentery, and back fixation suture to the mesentery (Figure 3I).

Postoperative observation

The drinking water was replaced with 5% dextrose-saline for the first 3 postoperative days. Penicillin (7.5 × 10⁴ IU/kg) was administered subcutaneously on postoperative days 1-3 to prevent infection. Additionally, the catheter was checked periodically for appropriate tube length and direction and patency of bile flow. Blockage of the silicone tube and excessive exposure of the exit site, hair, and skin to bile were prevented. Because SD rats can chew on the tubes, any evidence of biting was addressed promptly by adjusting the tube length and route. If the tube had been chewed through and retracted subcutaneously, it was promptly re-exteriorized under isoflurane anesthesia to avoid subcutaneous accumulation of bile.

Postoperative LPS injection

Starting on postoperative day 4, LPS was injected through the silicone tube once every 5 days on days 4, 9, 14, and 19. Based on prior experimental data and dose-ranging work, an LPS dose of 1 mg/kg at a concentration of 10 mg/mL was selected. LPS was dissolved in 0.9% normal saline by vortex mixing at 70 °C until fully dissolved.

The LPS injection dose was calculated based on the body weight of each SD rat and administered using a 100-µL microinjector. Because the catheter was approximately 18 cm long and had a capacity of approximately 20 µL, an additional 20 µL of LPS was aspirated in addition to the calculated dose to ensure delivery of the correct LPS dose into the bile duct, without an additional 20-µL injection of normal saline for flushing.

After each LPS injection, the end of the silicone tube was ligated with a 5-0 Mersilk suture. After 6 hours, the suture was released. Any LPS and bile overflowing from the end of the catheter upon release of the suture were cleaned away to avoid damage to the hair and skin.

Sampling

The necessary materials were prepared, including sterile disposable drapes, latex gloves, 50-mL centrifuge tubes, cryovials, a 2.5-mL syringe, phosphate-buffered saline, 4% paraformaldehyde, and the necessary surgical equipment. The rats were euthanized under isoflurane anesthesia, and the abdominal and thoracic cavities were immediately opened with surgical scissors. A 50-mL syringe equipped with a 21-G needle was inserted into the left ventricle, the right atrial appendage was incised, and perfusion with phosphate-buffered saline was performed. Successful perfusion was indicated when the liver became noticeably pale. The surrounding tissues were carefully dissected to identify the common bile duct, and bile was aspirated using a 2.5-mL syringe and placed in cryovials. Liver and biliary tissue samples were collected and preserved in 50-mL centrifuge tubes containing paraformaldehyde. The liver and bile duct tissues were then processed for paraffin embedding and sectioning, and the bile samples were subjected to smear analysis and FT-IR. Finally, the carcass of each SD rat was wrapped in a medical waste bag, appropriately sealed, and placed in a designated freezer for disposal.

RESULTS

To evaluate the feasibility and stability of this modeling method, 12 male SD rats were included: 6 rats received staged LPS injections, and 6 served as controls and received an equal volume of saline. Over the course of the experiment, 2 rats died: 1 control rat died on postoperative day 3 from bile peritonitis caused by bile leakage, and 1 experimental rat died after the day-9 LPS injection, likely related to the surgical stress of catheter replacement and progressively worsening cholangitis, giving an overall survival rate of 83%. Model success was comprehensively verified using multiple objective indicators, including bile smear examination, FT-IR, and histopathology, and clear criteria for successful model establishment were defined accordingly. All experimental animals that survived to the study endpoint successfully developed pigment (calcium bilirubinate) stones. The remaining rats underwent catheterization, drug administration, and sampling without incident, indicating that the model is generally feasible and has good survival and reproducibility.

The silicone tube, sleeve assembly, and a suitable guidewire were the main supplies used in this model. Silicone was selected because of its elasticity and good biocompatibility. Deposited bile can also adhere to the inner wall of the silicone tube to form a biofilm, gradually decreasing the effective luminal diameter—functionally equivalent to the gradual biliary constriction and poor open-ended biliary drainage of lithiasis-associated hepatobiliary cholangitis[14]. The average diameter of the common bile duct of SD rats ranges from 0.5 mm to 1.0 mm; hence, a silicone tube with inner and outer diameters of 0.3 mm and 0.8 mm, respectively, was chosen.

Bile collection and macroscopic assessment

Bile was collected on day 24. In experimental animals, approximately 1 mL of bile was aspirated by direct puncture of the dilated common bile duct with a 2.5-mL syringe. In controls, because bile drainage remained patent and the duct was not markedly dilated, less bile was obtained. Bile from the experimental group was darker in color and had increased viscosity, with visible flocculent material and sediment. Control bile was clear and transparent, with no obvious sediment.

Bile smear microscopy

Based on previously reported methods, bile was centrifuged at 12000 rpm for 10 minutes to obtain sediment[10]. After removal of the supernatant, the sediment was smeared and examined under a microscope. Yellow stone-like crystals were visible in the bile sediment of model-group rats (Figure 4A-D). In the control group, no obvious sediment formed after centrifugation; an equivalent volume of the centrifuged lower-layer bile was smeared, and no crystal-like material was observed (Figure 4E-H).

Figure 4 Microscopic bile smears. A-D: Experimental group (LPS-treated) bile sediment smears at increasing magnifications, showing clearly visible yellow stone-like crystal aggregates with increasing detail at higher magnification; E-H: Control group bile smears after centrifugation at magnifications of 4 ×, 10 ×, 20 ×, and 40 ×, respectively, showing no crystals; I-L: Smears of ground clinical hepatolithiasis stone specimens at the same magnifications, showing yellow crystal morphology essentially consistent with that of the LPS group. LPS: Lipopolysaccharide; NC: Negative control.

For direct comparison, clinically confirmed hepatolithiasis specimens (Ethics Approval No. 2025 L309) were dried in a 60 °C oven for 12 hours, fully ground, and sieved to obtain stone powder with a particle diameter of 0.05 mm. One gram of stone powder was mixed with 1 mL of normal saline and examined microscopically. The yellow crystal morphology in the SD rat model group was essentially consistent with that of the clinical specimens (Figure 4I-L).

FT-IR spectroscopic analysis

Bile sediment was dried in a 60 °C oven for 12 hours. The dried sediment was mixed with potassium bromide at a ratio of 1% (w/w), pressed into a semitransparent pellet, and analyzed using a Thermo Scientific NICOLET iS20 FT-IR spectrophotometer over the range of 400-4000 cm^-1 (resolution, 4 cm^-1; 50 scans per sample; absorbance mode)[15,16]. The same procedure was first applied to clinically confirmed hepatolithiasis specimens to validate the method; the spectrum obtained was consistent with the characteristic calcium bilirubinate spectrum reported in the literature (Figure 5A).

Figure 5 Fourier transform infrared spectroscopy. A: FT-IR spectrum of clinically confirmed calcium bilirubinate hepatolithiasis specimens. Key absorption bands: Broad peak at 3600-2400 cm^-1 (bound water); 3-peak combination at approximately 1637/1633/1630 cm^-1, approximately 1576 cm^-1, and approximately 1541 cm^-1 (characteristic calcium bilirubinate signature); C-H stretching at 2916/2849 cm^-1 (cholesterol). This spectrum serves as the reference standard confirming the analytical method; B: FT-IR spectrum of bile sediment from the experimental group (LPS-treated rats). The spectrum closely matches the clinical reference (Figure 5A): Broad bound-water absorption at 3600-2400 cm^-1; 3-peak combination near 1633/1630 cm^-1, 1599 cm^-1, and the 1500-1400 cm^-1 region, confirming calcium bilirubinate; C-H stretching at 2917/2849 cm^-1, indicating a minor cholesterol component. Taken together, the sediment is predominantly calcium bilirubinate with a small amount of cholesterol. FT-IR: Fourier transform infrared spectroscopy; LPS: Lipopolysaccharide.

The FT-IR spectrum of SD rat bile sediment showed the following features characteristic of calcium bilirubinate: A broad absorption peak in the 3600-2400 cm^-1 region (bound water) and a 3-peak combination near 1633/1630 cm^-1, 1599 cm^-1, and the 1500-1400 cm^-1 region, which is the hallmark signature of calcium bilirubinate. Cholesterol-related C-H stretching vibrations at approximately 2917/2849 cm^-1 were also present, indicating a minor cholesterol component (Figure 5B). Based on these features, the rat stone sediment was composed predominantly of calcium bilirubinate with a small amount of cholesterol.

Histopathological assessment

Pathological changes in hepatobiliary tissue were assessed according to the Scheuer scoring system[17]. On each hematoxylin- and eosin-stained section, no fewer than 5 representative portal tracts of varying sizes were randomly selected at × 20 magnification[18]. Inflammation (G0-G4) and fibrosis (S0-S4) were scored by an experienced pathologist who was blinded to the experimental design.

In the control group, the hepatic architecture was intact, the hepatic lobules were regularly arranged, and only sparse lymphocytes were present in the portal tracts, with no extension into the periportal parenchyma. No collagen deposition or fibrotic expansion was observed in any portal tract (Figure 6A and B). Inflammation scores were low overall (G0-G1), and all fibrosis scores were S0.

Figure 6 Hematoxylin and eosin staining of hepatobiliary tissue. A: Portal tract at 10 × magnification, showing intact hepatic lobule architecture with minimal lymphocytes in the portal tract, no periportal extension, and no fibrosis (control group); B: Portal tract at 20 × magnification, showing the inset region from A with normal portal structures, sparse inflammatory cells, and no collagen deposition (fibrosis score S0, inflammation score G0-G1; control group); C: Portal tract at 10 × magnification, showing a markedly expanded portal tract with dense inflammatory infiltrate, prominent bile duct reaction, and periportal inflammation; the inset highlights the area shown in D (experimental group; LPS-treated); D: Portal tract at 20 × magnification, showing dense inflammatory cell infiltration, bile duct proliferation, and periductal collagen fiber deposition indicating portal-periportal fibrosis (inflammation score G1-G2, fibrosis score S1; experimental group; LPS-treated). LPS: Lipopolysaccharide.

In the model group, varying degrees of hepatobiliary cholangitis-related pathological changes were present: Portal tract expansion, abundant inflammatory cell infiltration, and marked bile duct reaction (Figure 6C and D). In some areas, inflammatory cells extended into the periportal parenchyma, indicating active portal-periportal inflammation. Collagen fiber deposition and periportal fibrotic changes were present in and around the portal tracts. Inflammation scores (G1-G2) and fibrosis scores (S1) were higher than those in controls.

DISCUSSION

This experiment developed and optimized an SD rat model of hepatolithiasis using bile duct catheterization and a series of LPS injections. The model simulated the mechanism of inflammation-induced pigment stone formation: The experimental animals demonstrated characteristic biliary inflammation, calcium bilirubinate crystallization under the influence of β-G, and increased bile viscosity associated with mucin 5AC overexpression, pathological alterations that closely resemble those of human hepatolithiasis. More importantly, repeated administration via the catheters enabled LPS to act directly in the bile duct, simulating the recurrent cholangitis of human hepatolithiasis and replicating the dynamic process of inflammation-to-stone formation in vivo. This overcomes the inability of short-term or single-stimulation models to capture the dynamic disease course.

During the development of hepatolithiasis, biliary bacterial infection and the chronic inflammatory response associated with its metabolite LPS are considered an important pathological basis[1]. As an endotoxin of Gram-negative bacteria, LPS is chemically stable and can persist in the biliary system for a long time, continuously stimulating bile duct epithelial cells and activating inflammation-related signaling pathways as a typical pathogen-associated molecular pattern[6,19]. LPS also upregulates β-G expression[7]. β-G hydrolyzes conjugated bilirubin into unconjugated bilirubin and glucuronic acid; the unconjugated bilirubin then combines with calcium ions and precipitates, forming bile-insoluble calcium bilirubinate, which promotes pigment-type hepatolithiasis[7,8]. Mucins, high-molecular-weight glycoproteins secreted by the biliary epithelium, are induced and abnormally expressed via the protein kinase C/nuclear factor-kappa B/c-Myc signaling axis when LPS stimulates infected or inflamed bile duct epithelium, increasing bile viscosity and providing a favorable microenvironment for calcium bilirubinate deposition and stone formation, thereby further promoting hepatolithiasis[6,19,20]. Thus, hepatolithiasis is not caused by a single factor but results from the combined effects of LPS-mediated persistent biliary injury and coordinated regulation of β-G activity and mucin secretion.

Calcium bilirubinate was the major component of the stones, as confirmed by bile smear examination and FT-IR analysis of the samples. The presence of a minor cholesterol component indicates that this model can at least partially replicate the pathogenesis of mixed-component stones[15,16]. Hematoxylin and eosin staining revealed evidence of biliary epithelial damage, biliary fibrosis, and inflammatory cell infiltration. This model morphologically recapitulates biliary compositional changes, stone crystal growth, and biliary inflammatory responses and exhibits pathological changes at the molecular and histological levels comparable to those of clinical hepatolithiasis. The current model shows improved stability and reproducibility compared with dietary induction alone, LPS injection alone, or catheterization alone[9-12].

Several factors should be considered when developing a stable model of biliary stones. This study suggests that rats weighing at least 400 g are preferable because they have a sufficiently large common bile duct diameter and can be more easily manipulated; larger rats also better withstand chronic postoperative debilitation.

Compared with humans, the most notable anatomical difference in the SD rat biliary system is the absence of a gallbladder; thus, bile flows continuously into the duodenum without the storage and concentration processes that normally occur in the gallbladder[21]. In addition, the rat common bile duct is thinner and shorter, which places higher technical demands on catheterization. In this study, catheterization success was improved and the risk of catheter dislodgement was reduced by using a small-diameter silicone catheter, guidewire-assisted insertion, and multipoint fixation. On the other hand, although the rat liver differs from the human liver in lobar structure, it shows a high degree of biological similarity in bile duct epithelial structure, bile formation, and LPS-induced inflammatory responses. Notably, because SD rats lack a gallbladder, interference from gallbladder cholesterol stone formation is largely avoided, allowing the model to more closely reflect cholangitis-driven pigment stone formation and better align with the aim of studying primary intrahepatic bile duct stones.

In terms of LPS dosage, because there were no previous data on effective intraductal LPS doses in the literature, dose-ranging experiments were conducted at 5 mg/kg, 3 mg/kg, 2 mg/kg, 1 mg/kg, and 0.5 mg/kg, with LPS doses used in previous animal studies as a reference[22-24]. At high doses (greater than 2 mg/kg), acute hepatic failure and rapid deterioration occurred, making long-term observation impossible. At 0.5 mg/kg, stimulation was insufficient to maintain an inflammatory state in the biliary system. A dose of 1 mg/kg induced nonlethal biliary inflammation and was thus selected as the standard dose.

Concerning materials and methods, microsurgical instruments are highly recommended; precise handling greatly limits postoperative complications. Catheterization techniques and the catheter material used are important factors affecting model success and stability. Silicone was selected based on its biocompatibility, flexibility, and ability to support biofilm deposition on the inner wall, thereby recreating the gradual luminal constriction that occurs with chronic biliary inflammation[14].

Catheter dislodgement was avoided by the sleeve design. Multiple fixation sites and an appropriate insertion depth decrease the risk of dislodgement and biliary leakage, enhancing survival and the feasibility of the procedure. The subcutaneous routing of the external catheter, including the exit point, length outside the body, and orientation of the visible segment, is a vital part of the modeling process. Appropriate subcutaneous catheter length, exit site position, and external segment orientation help prevent the rat from biting through the tube, thereby avoiding failure to ligate the tube after drug delivery and preventing bile leakage or other potentially fatal complications.

Because the upper body is naturally contacted by the forelimbs in SD rats, the external catheter is prone to traction and biting. Constant observation of the catheter is thus required. Alterations in suture tension, external catheter length, or routing must be corrected as soon as possible to prevent dislodgement or subcutaneous accumulation of bile. Both rat deaths during modeling were associated with bile leakage and acute inflammatory exacerbation, indicating that future research should further improve the details of the procedure and its management.

Although the model has advantages, it also has limitations. The SD rat bile duct is very fine and requires highly specialized microsurgical skills; technical irregularities may affect the success and reproducibility of the model. Although the modeling period is sufficiently long (approximately 4 weeks) to induce stone formation, it is not long enough to completely recapitulate the years-long chronic evolution of human hepatolithiasis[1-3]. Individual variation in inflammatory responses and stone formation cannot be entirely eliminated, which may affect the use of this model for long-term disease studies and therapeutic evaluation. Caution should therefore be exercised when extrapolating the results to clinical settings.

The primary aim of this study was to establish and validate a stable, reproducible SD rat model of hepatolithiasis rather than to evaluate the therapeutic effect of a drug or other intervention. The focus was therefore on whether the model could be stably established and on the reproducibility of the methodology rather than on detecting small statistical differences between continuous variables. Regarding sample size, we strictly followed the 3R principles of animal research (replacement, reduction, and refinement), minimizing animal use while ensuring model reliability and adequate validation. Successful model establishment was determined using multiple objective indicators: Bile smear examination, FT-IR, and histology. All animals that survived to the study endpoint met the criteria for success, demonstrating good stability and consistency. We therefore consider the current sample size adequate for the purposes of model establishment and methodological validation. In future mechanistic or drug-intervention studies, a formal statistical power analysis can be performed based on specific study endpoints and expected effect sizes to determine a more appropriate sample size.

CONCLUSION

The SD rat hepatolithiasis model developed and optimized in the present study, which includes bile duct catheterization followed by staged LPS injections, simulates the pathological mechanism of hepatolithiasis and was validated by morphological, histological and spectroscopic analyses. It provides a viable research model for extending the observation period; enabling molecular-level analyses; investigating new interventions such as anti-inflammatory drugs, bile acid modulators, and β-G inhibitors; and studying the mechanisms of hepatolithiasis-related complications such as biliary fibrosis, cholestatic liver injury, and cholangiocarcinoma. With further refinement, this model may continue to play an important experimental role in bridging the gap between basic research and clinical translation in hepatolithiasis.

ACKNOWLEDGMENTS

We sincerely thank the researchers whose studies are referenced in this article and our co-authors for their valuable collaboration and support. We are also grateful to Professor Zhi-Gang Ma from the School of Chemical Sciences and Engineering at Yunnan University for his support of our research.