Extrahepatic Bile Duct and Gall Bladder Dissection in Nine-Day-Old Mouse Neonates

Summary

For the observation of murine neonatal bile duct disorders, an intact bile duct and efficient preparation are required. Therefore, a new approach for isolating the entire extrahepatic bile duct system in murine neonates was successfully developed while maintaining the integrity of the bile duct.

Abstract

The dissection of murine neonatal bile ducts has been described as difficult. The main aim of the described standard operating procedure is the isolation of the extrahepatic bile duct (EBD) in mouse neonates without damaging the bile duct during preparation. Because of its exceptionally close preparation compared to the cholangiocytes cell line and harvesting of the entire extrahepatic bile duct system (EBDS), the described approach is extremely useful in researching animal models of newborn bile duct disorders, such as biliary atresia. After euthanasia, the peritoneal cavity was accessed, and the bile duct system, duodenum, and liver were extracted with the unique En-bloc-Resection (EbR). The extracted sample is placed on a foam mat, and the EBD is dissected from contaminating cells atraumatically without necessary touch. The dissection of the entire EBDS is a significant advantage of this method. Caution must be taken due to the small size and amount of bile duct tissue. Using the described technique, there is no damage to the cholangiocytes. Further, the purity of the technique is reproducible (n = 10). Therefore, optimally comparable samples can be harvested. Furthermore, no bile duct tissue is harmed, because any contact with the bile duct system can be avoided during preparation, leaving the bile fluid inside the gall bladder. Most importantly, while performing the final gall bladder and bile duct dissection, atraumatic microinstruments were used only slightly lateral of the bile duct without squeezing it. This is the key to a clean and intact sample, and essential for further histological investigation or the isolation of cholangiocytes. To summarize, the described innovative dissection technique enables especially inexperienced operators with the necessary equipment to isolate the EBDS as cleanly as possible.

Introduction

The genesis and progression of cholangiopathies such as biliary atresia, primary sclerosing cholangitis (PSC), and primary biliary cholangitis (PBC) are either unknown or incomplete^1,2. The limited understanding of the origin and progression of those diseases leads to a paucity of therapy options³. The most difficult obstacle in studying neonatal bile duct disorders is gaining a molecular understanding of the pathophysiology. One of the essential keys to a better understanding of molecular pathology is the best possible observation of affected tissue. To avoid reduced comparability and discrepancies between research, such as observing potential viral etiology of biliary atresia⁴, the need for the best possible preparation and sharing of the performed dissection techniques arise. A pure preparation of the target tissue is necessary for later microscopical investigations or breeding cell- and 3D-organoid cultures. However, in murine neonatal disorders, tissue samples are rare and only occur in a small amount due to the very small size. Regarding bile duct disorders, difficulties in a clean preparation of bile ducts in murine neonates have been described⁵. Due to the neonatal stage of development, tissue differentiation is not overly advanced, which complicates preparation and increases the difficulty compared to the preparation of adult samples. Therefore, the operating workgroup investigated a novel strategy for preparing the EBDS in a neonatal mouse model. In the present study, the technique allows an efficientdissection of each sample.

The bile duct system is intraperitoneally placed in the right upper abdomen, arising from the liver. The gall bladder is located underneath the visceral surface of the liver's right lobe. The bile duct, together with the portal vein and hepatic artery, is embedded in the hepatoduodenal ligament. It joins the liver and the duodenum directly and drains bile fluid into the duodenum⁶. Anatomically, the bile duct is divided into the right and left hepatic ducts, the common hepatic duct, the cystic duct, and the Ductus choledochus, which is formed by the confluence of the cystic duct and the common hepatic duct⁷. This one eventually empties bile fluid and saliva from the pancreatic duct into the duodenum via the Ampulla of Vater.

Cholangiocytes line the bile duct intra- and extrahepatically, dwelling in a complicated anatomic niche where they assist in bile production and homeostasis⁸. Bile fluid passes these specialized epithelial cells in high concentrations daily. In particular, the HCO3- umbrella maintenance is very important for protecting against bile acid toxicity⁹. Cholangiocytes are the first line of defense in the hepatobiliary system against, for example, luminal microorganisms¹⁰. The cholangiocytes' defense efficacy against toxic assaults may be weakened by genetic predisposition. A toxic overload causes damage and destruction and can therefore lead to cholangiopathies. Furthermore, the developing bile duct is not completely capable of all self-protective mechanisms, leading to a higher susceptibility to environmental toxins in neonatal bile ducts¹¹.

Protocol

Following ethical approval (N045/2021), male and female C57BL/6 mice neonates were observed until 9 days old. The animals were born and provided for experimental purposes by the animal facility of the University Medical Center Hamburg-Eppendorf, Hamburg, Germany. The neonates were housed in a cage together with their parent animals. The environmental conditions were controlled in temperature (20-24 °C), 12:12 h light-dark cycle, and relative humidity of 40%-70%.

1. Experimental preparation

1. Prepare the required equipment for surgical operation, including scissors, forceps, etc. (see Table of Materials).
2. Place disinfected and autoclaved instruments on a sterile surface next to the operation table.
3. Euthanize a P9-aged neonatal mouse quickly by decapitation with an operating scissor. Place the body of the euthanized neonatal mouse on a sterile operation field. Dissect the EBDSs in 9-day-old mouse neonates (step 5).
   ​NOTE: The described dissection procedure gives any scientist the proper tools to remove the EBDS from neonatal and older mice. The older the mouse, the easier the preparation.

2. Access to the peritoneal cavity

1. Grasp the skin above the location of the urinary bladder with forceps. Incise a 2 mm diameter hole into the skin using scissors, without damaging the peritoneum and underlying structures. Expand the cut to the location of the decapitation, following the left front axillar line. Remove the skin from the left to the right side with the atraumatic forceps.
2. Take hold of the peritoneum surrounding the spleen. Gently lift it up until the peritoneum resembles a tent-like structure and cut a 1 mm diameter hole in the center. Wait for the "peritoneal tent" to fill up with air. Use a 10x microscopic magnification for this and the following steps.
3. Cut off the peritoneum in a window framed by the lower ribs, both lateral abdominal regions, and the lower bladder area to ensure full access to the liver, bile duct system, stomach, small intestines, and colon.
   ​NOTE: To improve access to the liver, an additional incision can be performed, removing the three lowest ribs while leaving the xiphoid process, falciform ligament, liver, and bile duct structures intact. The view of the liver is easy to obtain.

3. Examination of the gall bladder and bile ducts

NOTE: Ensure to keep the sample wet on a regular basis during all of the following steps.

1. Carefully pull the xiphoid process in a cranioventral position to examine the gall bladder.
   NOTE: The tension on the falciform ligament increases with this motion, and the attached gall bladder becomes visible.
   1. Perform only a slight pull to avoid uncontrollable tearing of the falciform ligament, which could lead to the gall bladder tearing from the bile duct system. Release the pull of the xiphoid process before the next step.
2. Gently pull down the duodenum to free the bile duct system.
   ​NOTE: As the tension on the hepatoduodenal ligament increases, the bile duct tissue becomes visible.

4. En-bloc-resection

1. Perform the lower en-bloc-mobilization following the steps below.
   1. Identify the duodenal papilla, which connects the bile duct system to the duodenum.
   2. Cut through the duodenum about 2 cm from to the right lateral side of the papilla.
   3. Cut through the pyloric area. Ensure that the stomach contents are present in the pyloric area between the cutting location and the duodenal papilla.
      NOTE: This is an important step for later security of orientation for the correct location of oral and aboral duodenal parts.
2. Perform the upper en-bloc-mobilization.
   1. Gently pull the xiphoid process and get access to the falciform ligament.
   2. Perform a 1 cm long cut through the falciform ligament, as close as possible to the xiphoid process, between the gall bladder and the xiphoid process. Ensure not to damage the gall bladder.
   3. Cut through the following connecting structures between the liver and the thorax: esophagus, inferior vena cava, thoracic aorta, all ligaments that surround the bare area of the liver, and all dorsally remaining tissue connections.
      ​NOTE: The en-bloc sample is completely dissected. It contains the liver, bile duct system, and the duodenal corpus, which is connected with the pyloric region of the stomach.

5. Final gall bladder and bile duct dissection

1. Perform the gross preparation following the steps below.
   1. Place the en-bloc sample on a foam pad usually used for dehydration. Use a 20x microscopic magnification and two microsurgical atraumatic forceps with a maximum tip size of 6 mm for this and the following steps.
   2. Assemble the sample on the foam pad. Reorganize the sample in the correct anatomical position. Flatten the oral and aboral part of the duodenum, performing a gentle movement.
   3. Start the movement at the duodenal papilla and continue to the cutting edges using atraumatic forceps. Smooth out the white pulpy contents of the stomach, which will only occur in the oral part of the duodenum. Ensure to identify the oral and the aboral part of the duodenum to rule out probable bile duct rotation.
   4. Cut away large remnants of hepatic tissue to begin dissection of the bile duct.
2. Perform the final isolation.
   1. Gently press the remaining hepatic tissue into the pores of the foam mat. Ensure that the scraping movements start from the bile duct system and lead to the liver boundaries.
   2. Transfer the sample to a cleaner position on the foam mat after a few scraping movements in various directions. Use the advantage of less squeezed liver tissue in the background to optimize the view for the best possible differentiation between EBDS and unwanted cells. Scrape the hepatic tissue from the bile duct until nothing, or as little as possible, of the hepatic tissue remains.
   3. Process the hepatoduodenal ligament until the isolated EBDS remains. Remove intraligamentous blood vessels like the hepatic artery, portal vein, and small remnants. Remove this delicate filament, with a gentle pull and high care, to the left lateral side. This dissection step might have already started unintentionally or partly completed during the prior removal of hepatic tissue, which in the end leads to the same result.
      NOTE: The blood vessels are emerging as a white and very delicate filament approximately 3-5 mm orally of the duodenal papilla and join the hepatoduodenal ligament from the left lateral side, accumulating with the bile duct to the glissonian triad. With the completion of this step, the final sample is completely isolated. If there is a need for a record, images can be captured after organizing the bile duct structures into their anatomical position (Figure 1). Doing the last preparation steps on a foam mat is recommended because the sample will not stick as much to the surface of the operation field. If the pores are wet, the bile duct levitates or floats. When moving the sample, it will not stick as firmly as it would with preparation on the operation cloth, resulting in no ripping while moving the sample.

6. Preparation for histological analysis

1. Put the isolated sample in a buffered solution, special medium, or formalin-containing fixatives (see Table of Materials) as soon as possible after the dissection.
2. Chose a suitable storage solution depending on further planned processing steps.
   CAUTION: Use formalin-containing fixatives only under an air vent because of acute toxicity, corrosivity, and diverse health hazards.
   NOTE: In the presented study, the EBDS samples have been inserted in paraformaldehyde, dehydrated, and embedded in paraffin. They have been stored at room temperature and cooled down prior to sectioning. In a warming cabinet, 2 µm slices were kept overnight and stained using conventional hematoxylin and eosin⁴ (see Table of Materials).

Results

Figure 1A shows the EBDS of a murine neonate, which was dissected with the described technique. Microscopically, no further hepatic tissue is visible. The hepatic tissue has been removed during the final isolation steps of the protocol and could easily be distinguished from bile duct tissue regarding color and consistency. Figure 1B displays the isolated sample compared to a millimeter scale. The EBD's length (measured from gall bladder to duodenal papilla) is l...

Discussion

This article reported and discussed the creation and validation of a new surgical approach for removing the EBDS of euthanized neonatal mice. Microscopical and histological findings reveal that the approach quickly detects EBDs and dissects them near the duct's margins, even in neonatal mice. Only surgical instruments and a microscope with a 20x magnification are required for the described protocol. Furthermore, the approach allows the isolation of the entire EBDS. The technique is highly efficient, straightforward, ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
2-Propanol	CHEMSOLUTE	11365000	used as a dehydrating agent
30 G canula	B Braun/Sterican, Melsungen Germany	4656300	canula for hydration of the sample
Air vent	C + P Möbelsysteme GmbH & Co. KG, Breidenbach, Germany	Tec-Ononmic AZ 1200	the use of an air vent helps to avoid inhalation of formalin-containing fixatives
Aqua ad injectabilia Braun	B Braun, Melsungen, Germany	2351744	saline; Container: Mini-Plasco connect, 20 x 10 mL, sterile
Bigger microsurgical Forceps	DIADUST von Aesculap, Trossingen Deutschland	FD253R	straight, 180 mm (7"), platform tip, round handle, width: 0,800 mm, diamond dust coated, non-sterile, reusable optional tool for observation and every step of preparation except very final preparation; Dividing skin of the peritoneum
Camera “SmartCAM 5”	Basler and Vision Engineering, Send, United Kingdom	EVC131A	optional Lynx Exo camera modul: sensortype: CMOS, resolution 2560 x 1920 pixels, sensor size: 1/2"; Used for videoproduction and technical evaluation
Dehydration machine/Citadel 2000 Tissue Processor	Fisher Scientific GmbH, Schwerte, Germany	12612613	used for automatic dehydration, short program (approx. 4.8 h)
Dehydration sponge	Carl Roth, Karlsruhe, Germany	TT56.1	sponge for final dissection step, other sponges/foam pads with a minimum pore size of 60 pores per inch are also suitable, the use of  two foam pads per embedding cassette is recomended to cover the sample from below and above to prevent sliding through the perforation of the embedding cassettes
Dulbecco´s Phosphat Buffered Saline (PBS)	Gibco	14190-144	Doesn´t contain Calzium or Magnesium, 500 mL
Embedding cassettes	Engelbrecht GmbH, Edermünde, Germany	17990
Eosin	MEDITE Medical GmbH, Burgdorf, Germany	41-6660-00	staining solution, ready to use
Fine Scissors CeramaCut	FST, Heidelberg Germany	14959-09	Tips: Sharp-Sharp, Alloy / Material: Ceramic Coated Stainless Steel, Serrated:, Yes; Feature: CeramaCut, Tip Shape: Straight, Cutting Edge: 22 mm, Length: 9 cm; Skin incision, incision of the peritoneal window
Graefe Forceps	FST, Heidelberg Germany	11051-10	Length: 10 cm, Tip Shape: curved, serrated, Tip width: 0.8 mm, Tip Dimensions: 0.8 x 0.7 mm, Alloy /Material: Stainless Steel
Hematoxylin	MEDITE Medical GmbH, Burgdorf, Germany	41-5130-00	staining solution, ready to use
Highresolotion microscope	Vision Engineering, Send United Kingdom	EVO503	Capable of enlargement up to 60x magnification, only 6x to 20x magnification were used
Microscope	Olympus Optical CO, Ltd., Hamburg, Germany	BX60F5
Microscope Cover Glases	Marienfeld, Lauda-Königshofen, Germany	101244	60 mm broad, made of SCHOTT D 263 glass
Microscope Slides	R. Langenbrinck GmbH, Emmendingen, Germany	03-0060
Microtome	Leica, Nußloch, Germany	SM2010R	Tool for sectioning (2 µm-slices)
Omnifix-F 1 mL syringe	B Braun, Melsungen, Germany	9161406V	syringe without canula
Paraffin	Sakura Finetec, Torrance, USA	4511	Tissue-Tek Paraffin Wax Tek III, without DMSO
Paraffin embedding machine	MEDITE Medical GmbH, Burgdorf, Germany	TES 99	The embedding machine used in this study contained the following three individual modules: TES 99.420, TES 99.250, TES 99.600. The sample should be embedded in Paraffin directly after the dehydration, no interim storage in a fridge should be performed due to possible shrinking and moisture in the fridge
Paraformaldehyde (PFA)	Morphisto	1176201000	Prepare 1 mL Aliquots in 2 mL Eppendorf conical Tubes for liver samples and 0.5 mL Aliquots in 1 mL Eppendorf conical Tubes for extrahepatic bile duct samples, 4% in PBS ph 7.4
Small Microsurgical Forceps	EPM (Erich Pfitzer Medizintechnik), Bütthard, Bayern, Germany	(00)165	Round handle, straight, 0.3 mm tip, tool for observation and every step of preparation, especially useful in final preparation
Stainless Steel Ruler	Agntho's AB, Lidingö, Sweden	30085-15	150mm With Metric & Inch Graduations
Surgical Scissors – Sharp-blunt for decapitation	FST, Heidelberg Germany	14001-14	Device for decapitation
Warming cabinet	Haraeus, Hanau, Germany	T 6060	the sliced samples should be kept in the warming cabinet to ensure the attachement of the sample on the microscope slides