The authors acknowledge Johanna Hagens, Pauline Schuppert, Clara Philippi, PD Dr. med Christian Tomuschat, Svenja Warnke, PD Dr. Diana Lindner, Prof. Dr. Dirk Westermann, Miriam Tomczak, Nicole L&#252;der, Nadine Kurzawa, Dr. rer nat. Laia Pagerols Raluy, Birgit Appl, and Magdalena Trochimiuk for their contributions.&#160;Hans Christian Schmidt was financially supported by the Else Kr&#246;ner-Fresenius-Stiftung iPRIME Scholarship (2021_EKPK.10), UKE, Hamburg.

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 &#176;C), 12:12 h light-dark cycle, and relative humidity of 40%-70%.
1. Experimental preparation
<ol>
	<li>Prepare the required equipment for surgical operation, including scissors, forceps, etc. (see Table of Materials).</li>
	<li>Place disinfected and autoclaved instruments on a sterile surface next to the operation table.</li>
	<li>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). 
	&#8203;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.</li>
</ol>
2. Access to the peritoneal cavity
<ol>
	<li>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.</li>
	<li>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 &#34;peritoneal tent&#34; to fill up with air. Use a 10x microscopic magnification for this and the following steps.</li>
	<li>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. 
	&#8203;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.</li>
</ol>
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.
<ol>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Gently pull down the duodenum to free the bile duct system. 
	&#8203;NOTE: As the tension on the hepatoduodenal ligament increases, the bile duct tissue becomes visible.</li>
</ol>
4. En-bloc-resection
<ol>
	<li>Perform the lower en-bloc-mobilization following the steps below.
	<ol>
		<li>Identify the duodenal papilla, which connects the bile duct system to the duodenum.</li>
		<li>Cut through the duodenum about 2 cm from to the right lateral side&#160;of the papilla.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Perform the upper en-bloc-mobilization.
	<ol>
		<li>Gently pull the xiphoid process and get access to the falciform ligament.</li>
		<li>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.</li>
		<li>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. 
		&#8203;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.</li>
	</ol>
	</li>
</ol>
5. Final gall bladder and bile duct dissection
<ol>
	<li>Perform the gross preparation following the steps below.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Cut away large remnants of hepatic tissue to begin dissection of the bile duct.</li>
	</ol>
	</li>
	<li>Perform the final isolation.
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
6. Preparation for histological analysis
<ol>
	<li>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.</li>
	<li>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 &#181;m slices were kept overnight and stained using conventional hematoxylin and eosin4 (see Table of Materials).</li>
</ol>

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	<li>Liwinski, T., Schramm, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prim&#228;r+sklerosierende+Cholangitis.">Prim&#228;r sklerosierende Cholangitis.</a> Der Internist. 59 (6), 551-559 (2018).</li><li>Kobayashi, H., Stringer, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biliary+atresia.">Biliary atresia.</a> Seminars in Neonatology. 8 (5), 383-391 (2003).</li><li>Patman, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biliary+tract:+Newly+identified+biliatresone+causes+biliary+atresia.">Biliary tract: Newly identified biliatresone causes biliary atresia.</a> Nature Reviews Gastroenterology &amp; Hepatology. 12 (7), 369 (2015).</li><li>Mack, C. L., Sokol, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unraveling+the+pathogenesis+and+etiology+of+biliary+atresia.">Unraveling the pathogenesis and etiology of biliary atresia.</a> Pediatric Research. 57 (5), 87-94 (2005).</li><li>Karjoo, S., Wells, R. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+neonatal+extrahepatic+cholangiocytes.">Isolation of neonatal extrahepatic cholangiocytes.</a> Journal of Visualized Experiments. (88), e51621 (2014).</li><li>Strazzabosco, M., Fabris, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+anatomy+of+normal+bile+ducts.">Functional anatomy of normal bile ducts.</a> The Anatomical Record: Advances in Integrative Anatomy and Evolutionary Biology. 291 (6), 653-660 (2008).</li><li>Nakanuma, Y., Hoso, M., Sanzen, T., Sasaki, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microstructure+and+development+of+the+normal+and+pathologic+biliary+tract+in+humans,+including+blood+supply.">Microstructure and development of the normal and pathologic biliary tract in humans, including blood supply.</a> Microscopy Research and Technique. 38 (6), 552-570 (1997).</li><li>Banales, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cholangiocyte+pathobiology.">Cholangiocyte pathobiology.</a> Nature Reviews Gastroenterology &amp; Hepatology. 16 (5), 269-281 (2019).</li><li>de Buy Wenniger, L. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+cholangiocyte+glycocalyx+stabilizes+the+'biliary+HCO3-+umbrella':+an+integrated+line+of+defense+against+toxic+bile+acids.">The cholangiocyte glycocalyx stabilizes the 'biliary HCO3- umbrella': an integrated line of defense against toxic bile acids.</a> Digestive Diseases. 33 (3), 397-407 (2015).</li><li>Pinto, C., Giordano, D. M., Maroni, L., Marzioni, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+inflammation+and+proinflammatory+cytokines+in+cholangiocyte+pathophysiology.">Role of inflammation and proinflammatory cytokines in cholangiocyte pathophysiology.</a> Biochimica et Biophysica Acta (BBA)-Molecular Basis of Disease. 1864 (4), 1270-1278 (2018).</li><li>Khandekar, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordinated+development+of+the+mouse+extrahepatic+bile+duct:+Implications+for+neonatal+susceptibility+to+biliary+injury.">Coordinated development of the mouse extrahepatic bile duct: Implications for neonatal susceptibility to biliary injury.</a> Journal of Hepatology. 72 (1), 135-145 (2020).</li><li>Grundmann, D., Klotz, M., Rabe, H., Glanemann, M., Sch&#228;fer, K. -. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+high-purity+myenteric+plexus+from+adult+human+and+mouse+gastrointestinal+tract.">Isolation of high-purity myenteric plexus from adult human and mouse gastrointestinal tract.</a> Scientific Reports. 5 (1), 9226 (2015).</li><li>Ishii, M., Vroman, B., LaRusso, N. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+morphologic+characterization+of+bile+duct+epithelial+cells+from+normal+rat+liver.">Isolation and morphologic characterization of bile duct epithelial cells from normal rat liver.</a> Gastroenterology. 97 (5), 1236-1247 (1989).</li><li>Kumar, U., Jordan, T. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+culture+of+biliary+epithelial+cells+from+the+biliary+tract+fraction+of+normal+rats.">Isolation and culture of biliary epithelial cells from the biliary tract fraction of normal rats.</a> Liver. 6 (6), 369-378 (1986).</li><li>Vroman, B., LaRusso, N. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+characterization+of+polarized+primary+cultures+of+rat+intrahepatic+bile+duct+epithelial+cells.">Development and characterization of polarized primary cultures of rat intrahepatic bile duct epithelial cells.</a> Laboratory Investigation. 74 (1), 303-313 (1996).</li><li>Paradis, K., Sharp, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+duct-like+structure+formation+after+isolation+of+bile+ductular+cells+from+a+murine+model.">In vitro duct-like structure formation after isolation of bile ductular cells from a murine model.</a> Journal of Laboratory and Clinical Medicine. 113 (6), 689-694 (1989).</li></ol>

The authors declare no conflict of interest.

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&#39;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, ...

The genesis and progression of cholangiopathies such as biliary atresia, primary sclerosing cholangitis (PSC), and primary biliary cholangitis (PBC) are either unknown or incomplete1,2. 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,&#160;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&#39;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&#39; 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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-Propanol</td>
			<td>CHEMSOLUTE</td>
			<td>11365000</td>
			<td>used as a dehydrating agent</td>
		</tr>
		<tr>
			<td>30 G canula</td>
			<td>B Braun/Sterican, Melsungen Germany</td>
			<td>4656300</td>
			<td>canula for hydration of the sample</td>
		</tr>
		<tr>
			<td>Air vent</td>
			<td>C + P M&#246;belsysteme GmbH &#38; Co. KG, Breidenbach, Germany</td>
			<td>Tec-Ononmic AZ 1200</td>
			<td>the use of an air vent helps to avoid inhalation of formalin-containing fixatives</td>
		</tr>
		<tr>
			<td>Aqua ad injectabilia Braun</td>
			<td>B Braun, Melsungen, Germany</td>
			<td>2351744</td>
			<td>saline; Container: Mini-Plasco connect, 20 x 10 mL, sterile</td>
		</tr>
		<tr>
			<td>Bigger microsurgical Forceps</td>
			<td>DIADUST von Aesculap, Trossingen Deutschland</td>
			<td>FD253R</td>
			<td>straight, 180 mm (7&#34;), 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</td>
		</tr>
		<tr>
			<td>Camera &#8220;SmartCAM 5&#8221;&#160;</td>
			<td>Basler and Vision Engineering, Send, United Kingdom</td>
			<td>EVC131A</td>
			<td>optional Lynx Exo camera modul: sensortype: CMOS, resolution 2560 x 1920 pixels, sensor size: 1/2&#34;; Used for videoproduction and technical evaluation</td>
		</tr>
		<tr>
			<td>Dehydration machine/Citadel 2000 Tissue Processor</td>
			<td>Fisher Scientific GmbH, Schwerte, Germany</td>
			<td>12612613</td>
			<td>used for automatic dehydration, short program (approx. 4.8 h)</td>
		</tr>
		<tr>
			<td>Dehydration sponge&#160;</td>
			<td>Carl Roth, Karlsruhe, Germany</td>
			<td>TT56.1</td>
			<td>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&#160; 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</td>
		</tr>
		<tr>
			<td>Dulbecco&#180;s Phosphat Buffered Saline (PBS)</td>
			<td>Gibco</td>
			<td>14190-144</td>
			<td>Doesn&#180;t contain Calzium or Magnesium, 500 mL</td>
		</tr>
		<tr>
			<td>Embedding cassettes</td>
			<td>Engelbrecht GmbH, Ederm&#252;nde, Germany</td>
			<td>17990</td>
			<td></td>
		</tr>
		<tr>
			<td>Eosin</td>
			<td>MEDITE Medical GmbH, Burgdorf, Germany</td>
			<td>41-6660-00</td>
			<td>staining solution, ready to use</td>
		</tr>
		<tr>
			<td>Fine Scissors CeramaCut</td>
			<td>FST, Heidelberg Germany</td>
			<td>14959-09</td>
			<td>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</td>
		</tr>
		<tr>
			<td>Graefe Forceps</td>
			<td>FST, Heidelberg Germany</td>
			<td>11051-10</td>
			<td>Length: 10 cm, Tip Shape: curved, serrated, Tip width: 0.8 mm, Tip Dimensions: 0.8 x 0.7 mm, Alloy /Material: Stainless Steel</td>
		</tr>
		<tr>
			<td>Hematoxylin</td>
			<td>MEDITE Medical GmbH, Burgdorf, Germany</td>
			<td>41-5130-00</td>
			<td>staining solution, ready to use</td>
		</tr>
		<tr>
			<td>Highresolotion microscope</td>
			<td>Vision Engineering, Send United Kingdom</td>
			<td>EVO503&#160;</td>
			<td>Capable of enlargement up to 60x magnification, only 6x to 20x magnification were used&#160;</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Olympus Optical CO, Ltd., Hamburg, Germany</td>
			<td>BX60F5</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Cover Glases</td>
			<td>Marienfeld, Lauda-K&#246;nigshofen, Germany</td>
			<td>101244</td>
			<td>60 mm broad, made of SCHOTT D 263 glass</td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>R. Langenbrinck GmbH, Emmendingen, Germany</td>
			<td>03-0060</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtome</td>
			<td>Leica, Nu&#223;loch, Germany</td>
			<td>SM2010R</td>
			<td>Tool for sectioning (2 &#181;m-slices)&#160;</td>
		</tr>
		<tr>
			<td>Omnifix-F 1 mL syringe</td>
			<td>B Braun, Melsungen, Germany</td>
			<td>9161406V</td>
			<td>syringe without canula</td>
		</tr>
		<tr>
			<td>Paraffin</td>
			<td>Sakura Finetec, Torrance, USA</td>
			<td>4511</td>
			<td>Tissue-Tek Paraffin Wax Tek III, without DMSO</td>
		</tr>
		<tr>
			<td>Paraffin embedding machine</td>
			<td>MEDITE Medical GmbH, Burgdorf, Germany</td>
			<td>TES 99</td>
			<td>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</td>
		</tr>
		<tr>
			<td>Paraformaldehyde (PFA)</td>
			<td>Morphisto</td>
			<td>1176201000</td>
			<td>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&#160;</td>
		</tr>
		<tr>
			<td>Small Microsurgical Forceps&#160;</td>
			<td>EPM (Erich Pfitzer Medizintechnik), B&#252;tthard, Bayern, Germany</td>
			<td>(00)165</td>
			<td>Round handle, straight, 0.3 mm tip, tool for observation and every step of preparation, especially useful in final preparation</td>
		</tr>
		<tr>
			<td>Stainless Steel Ruler</td>
			<td>Agntho&#39;s AB, Liding&#246;, Sweden</td>
			<td>30085-15</td>
			<td>150mm With Metric &#38; Inch Graduations</td>
		</tr>
		<tr>
			<td>Surgical Scissors &#8211; Sharp-blunt for decapitation</td>
			<td>FST, Heidelberg Germany</td>
			<td>14001-14</td>
			<td>Device for decapitation</td>
		</tr>
		<tr>
			<td>Warming cabinet</td>
			<td>Haraeus, Hanau, Germany</td>
			<td>T 6060</td>
			<td>the sliced samples should be kept in the warming cabinet to ensure the attachement of the sample on the microscope slides</td>
		</tr>
	</tbody>
</table>

extrahepatic bile duct and gall bladder dissection in nine-day-old mouse neonates

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.

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&#39;s length (measured from gall bladder to duodenal papilla) ...

Watch this Scientific Journal Video about Extrahepatic Bile Duct and Gall Bladder Dissection in Nine-Day-Old Mouse Neonates at JoVE.com

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

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.

The dissection of murine neonatal bile ducts has been described as difficult. The main aim of the described standard operating procedure is the isolation ...

extrahepatic-bile-duct-gall-bladder-dissection-nine-day-old-mouse

Research

JoVE Journal

Medicine

1.8K Views.  University Medical Center Hamburg-Eppendorf. 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.

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

Mouse Bladder Wall Injection

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.

Mouse bladder wall injection is a useful approach to orthotopically study bladder stem cell and cancer biology. This delicate microsurgical method can be mastered with careful technique and practice.

Mouse bladder wall injection is a useful technique to orthotopically study bladder phenomena, including stem cell, smooth muscle, and cancer biology. ...

An Orthotopic Model of Murine Bladder Cancer

In this straightforward procedure, bladder tumors are established in female C57 mice through the use of catheterization, local cauterization, and subsequent cell adhesion. After their bladders are transurethrally catheterized and drained, animals are again catheterized to permit insertion of a platinum wire into bladders without damaging the urethra or bladder. The catheters are made of Teflon to serve as an insulator for the wire, which will conduct electrical current into the bladder to create a burn injury. An electrocautery unit is used to deliver 2.5W to the exposed end of the wire, burning away extracellular layers and providing attachment sites for carcinoma cells that are delivered in suspension to the bladder through a subsequent catheterization. Cells remain in the bladder for 90 minutes, after which the catheters are removed and the bladders allowed to drain naturally. The development of tumor is monitored via ultrasound. Specific attention is paid to the catheterization technique in the accompanying video.

Bladder tumors are established in female mice in a minimally invasive fashion through catheterization, local cauterization, and subsequent adhesion of carcinoma cells to the burn sites.

In this straightforward procedure, bladder tumors are established in female C57 mice through the use of catheterization, local cauterization, and ...

Cannulation of the Mouse Submandibular Salivary Gland via the Wharton's Duct

Severe salivary gland hypofunction is frequently found in patients with Sj&ouml;gren's syndrome and those who receiving therapeutic 
irradiation in their head and neck regions for cancer treatment. Both groups of patients experience symptoms such as xerostomia (dry mouth), dysphagia 
(impaired chewing and swallowing), severe dental caries, altered taste, oro-pharyngeal infections (candidiasis), mucositis, pain and discomfort.
One innovative approach of regenerative medicine for the treatment of salivary gland hypo-function is speculated in RS Redman, E Mezey 
et al. 2009: stem cells can be directly deposited by cannulation into the gland as a potent method in reviving the functions of the impaired organ. Presumably, 
the migrated foreign stem cells will differentiate into glandular cells to function as part of the host salivary gland. Also, this cannulation technique is an 
expedient and effective delivery method for clinical gene transfer application.
Here we illustrate the steps involved in performing the cannulation procedure on the mouse submandibular salivary gland via the Wharton's duct 
(Fig 1). C3H mice (Charles River, Montreal, QC, Canada) are used for this experiment, which have been kept under clean conventional conditions at 
the McGill University animal resource center. All experiments have been approved by the University Animal Care Committee and were in accordance with the 
guidelines of the Canadian Council on Animal Care.
For this experiment, a trypan blue solution is infused into the gland through the opening of the Wharton's duct using a 
insulin syringe with a 29-gauge needle encased inside a polyethylene tube. Subsequently, the mouse is dissected to show that the infusions migrated 
into the gland successfully.

Cannulation of the Mouse Submandibular Salivary Gland via the Wharton&#039;s Duct

A protocol for the cannulation of the mouse submandibular salivary gland via the Wharton's duct is described. For this experiment, the trypan blue solution is used as a dyer to demonstrate how this technique effectively delivers infusions into the targeted gland, and to suggest the reliability of this new approach as a potential clinical drug/cell therapy for the regeneration of salivary glands.

Severe salivary gland hypofunction is frequently found in patients with Sj&ouml;gren's syndrome and those who receiving therapeutic 
irradiation in their ...

3-Dimensional Resin Casting and Imaging of Mouse Portal Vein or Intrahepatic Bile Duct System

In organs, the correct architecture of vascular and ductal structures is indispensable for proper physiological function, and the formation and maintenance of these structures is a highly regulated process. The analysis of these complex, 3-dimensional structures has greatly depended on either 2-dimensional examination in section or on dye injection studies. These techniques, however, are not able to provide a complete and quantifiable representation of the ductal or vascular structures they are intended to elucidate. Alternatively, the nature of 3-dimensional plastic resin casts generates a permanent snapshot of the system and is a novel and widely useful technique for visualizing and quantifying 3-dimensional structures and networks.
A crucial advantage of the resin casting system is the ability to determine the intact and connected, or communicating, structure of a blood vessel or duct. The structure of vascular and ductal networks are crucial for organ function, and this technique has the potential to aid study of vascular and ductal networks in several ways. Resin casting may be used to analyze normal morphology and functional architecture of a luminal structure, identify developmental morphogenetic changes, and uncover morphological differences in tissue architecture between normal and disease states. Previous work has utilized resin casting to study, for example, architectural and functional defects within the mouse intrahepatic bile duct system that were not reflected in 2-dimensional analysis of the structure1,2, alterations in brain vasculature of a Alzheimer's disease mouse model3, portal vein abnormalities in portal hypertensive and cirrhotic mice4, developmental steps in rat lymphatic maturation between immature and adult lungs5, immediate microvascular changes in the rat liver, pancreas, and kidney in response in to chemical injury6.
Here we present a method of generating a 3-dimensional resin cast of a mouse vascular or ductal network, focusing specifically on the portal vein and intrahepatic bile duct. These casts can be visualized by clearing or macerating the tissue and can then be analyzed. This technique can be applied to virtually any vascular or ductal system and would be directly applicable to any study inquiring into the development, function, maintenance, or injury of a 3-dimensional ductal or vascular structure.

A method of visualizing and quantifying the 3-dimensional structure of mouse hepatic portal vein or intrahepatic bile duct is described. This resin cast technique can also be applied to other ductal or vascular systems and allows for in situ visualization or quantification of a system's intact communicating architecture.

In  organs, the correct architecture of vascular and ductal structures is  indispensable for proper physiological function, and the formation and  ...

Urinary Bladder Distention Evoked Visceromotor Responses as a Model for Bladder Pain in Mice

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition characterized by increased urgency and frequency of urination, as well as nocturia and general pelvic pain, especially upon bladder filling or voiding. Despite years of research, the cause of IC/BPS remains elusive and treatment strategies are unable to provide complete relief to patients. In order to study nervous system contributions to the condition, many animal models have been developed to mimic the pain and symptoms associated with IC/BPS. One such murine model is urinary bladder distension (UBD). In this model, compressed air of a specific pressure is delivered to the bladder of a lightly anesthetized animal over a set period of time. Throughout the procedure, wires in the superior oblique abdominal muscles record electrical activity from the muscle. This activity is known as the visceromotor response (VMR) and is a reliable and reproducible measure of nociception. Here, we describe the steps necessary to perform this technique in mice including surgical manipulations, physiological recording, and data analysis. With the use of this model, the coordination between primary sensory neurons, spinal cord secondary afferents, and higher central nervous system areas involved in bladder pain can be unraveled. This basic science knowledge can then be clinically translated to treat patients suffering from IC/BPS.

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition characterized in part by pelvic pain. In order to study nervous system contributions to the condition, a physiological model of urinary bladder pain is used in mice and rats.

Approximately 3-8 million people in the United States suffer from interstitial cystitis/bladder pain syndrome (IC/BPS), a debilitating condition ...

State of the Art Cranial Ultrasound Imaging in Neonates

Cranial ultrasound (CUS) is a reputable tool for brain imaging in critically ill neonates. It is safe, relatively cheap and easy to use, even when a patient is unstable. In addition it is radiation-free and allows serial imaging. CUS possibilities have steadily expanded. However, in many neonatal intensive care units, these possibilities are not optimally used. We present a comprehensive approach for neonatal CUS, focusing on optimal settings, different probes, multiple acoustic windows and Doppler techniques. This approach is suited for both routine clinical practice and research purposes. In a live demonstration, we show how this technique is performed in the neonatal intensive care unit. Using optimal settings and probes allows for better imaging quality and improves the diagnostic value of CUS in experienced hands. Traditionally, images are obtained through the anterior fontanel. Use of supplemental acoustic windows (lambdoid, mastoid, and lateral fontanels) improves detection of brain injury. Adding Doppler studies allows screening of patency of large intracranial arteries and veins. Flow velocities and indices can be obtained. Doppler CUS offers the possibility of detecting cerebral sinovenous thrombosis at an early stage, creating a window for therapeutic intervention prior to thrombosis-induced tissue damage. Equipment, data storage and safety aspects are also addressed.

Cranial ultrasound (CUS) is a valuable tool for brain imaging in critically ill neonates. This video shows a comprehensive approach for neonatal (Doppler) CUS for both clinical and research purposes, including a bedside demonstration of the technique.

Cranial ultrasound (CUS) is a reputable tool for brain imaging in critically ill neonates. It is safe, relatively cheap and easy to use, even when a ...

Bile Duct Ligation in Mice: Induction of Inflammatory Liver Injury and Fibrosis by Obstructive Cholestasis

In most vertebrates, the liver produces bile that is necessary to emulsify absorbed fats and enable the digestion of lipids in the small intestine as well as to excrete bilirubin and other metabolic products. In the liver, the experimental obstruction of the extrahepatic biliary system initiates a complex cascade of pathological events that leads to cholestasis and inflammation resulting in a strong fibrotic reaction originating from the periportal fields. Therefore, surgical ligation of the common bile duct has become the most commonly used model to induce obstructive cholestatic injury in rodents and to study the molecular and cellular events that underlie these pathophysiological mechanisms induced by inappropriate bile flow. In recent years, different surgical techniques have been described that either allow reconnection or reanastomosis after bile duct ligation (BDL), e.g., partial BDL, or other microsurgical methods for specific research questions. However, the most frequently used model is the complete obstruction of the common bile duct that induces a strong fibrotic response after 21 to 28 days. The mortality rate can be high due to infectious complications or technical inaccuracies. Here we provide a detailed surgical procedure for the BDL model in mice that induce a highly reproducible fibrotic response in accordance to the 3R rule for animal welfare postulated by Russel and Burch in 1959.

Disruption of bile flow results in severe inflammatory cholestatic liver injury with a characteristic time-dependent sequence of morphological alterations. Here we present a protocol for the surgical ligation of the common bile duct in mice that allows to induce a strong fibrotic response after 21 to 28 days.

In most vertebrates, the liver produces bile that is necessary to emulsify absorbed fats and enable the digestion of lipids in the small intestine as well ...

Transcutaneous Microcirculatory Imaging in Preterm Neonates

Microcirculatory imaging (MI) is a relatively new research tool mainly used in the intensive care setting. MI provides a clear view of the smallest capillaries, arterioles and venules. The magnifying effect visualizes the flow pattern of erythrocytes through these vessels.
It's non-invasive character makes it suitable to apply in (preterm) neonates, even in cardiorespiratory unstable patients. In adults and children, MI is mainly performed sublingually, but this is not possible in preterm infants as these cannot cooperate and the size of the probe is problematic. In preterm infants, MI is therefore performed transcutaneously. Their thin skin makes it possible to obtain high quality images of peripheral microcirculation. 
In this manuscript we will demonstrate the method of transcutaneous MI in preterm infants. We will focus on the different techniques and provide tips to optimize image quality. The highlights of software settings, safety and offline analysis are also addressed.

Microcirculatory imaging (MI) is used to monitor peripheral perfusion in critically ill or preterm neonates. This manuscript and video demonstrates the optimal approach for obtaining high-quality images.

Microcirculatory imaging (MI) is a relatively new research tool mainly used in the intensive care setting. MI provides a clear view of the smallest ...

Isolation and Profiling of MicroRNA-containing Exosomes from Human Bile

Exosome research in the last three years has greatly extended the scope towards identification and characterization of biomarkers and their therapeutic uses.
Exosomes have recently been shown to contain microRNAs (miRs). MiRs themselves have arisen as valuable biomarkers for diagnostic purposes. As specimen collection in clinics and hospitals is quite variable, miRNA isolation from whole bile varies substantially. To achieve robust, accurate and reproducible miRNA profiles from collected bile samples in a simple manner required the development of a high-quality protocol to isolate and characterize exosomes from bile. The method requires several centrifugations and a filtration step with a final ultracentrifugation step to pellet the isolated exosomes. Electron microscopy, Western blots, flow cytometry and multi-parameter nanoparticle optical analysis, where available, are crucial characterization steps to validate the quality of the exosomes. For the isolation of miRNA from these exosomes, spiking the lysate with a non-specific, synthetic miRNA from a species like Caenorhabditis elegans, i.e., Cel-miR-39, is important for normalization of RNA extraction efficiency. The isolation of exosome from bile fluid following this method allows the successful miRNA profiling from bile samples stored for several years at -80 &#176;C.

Bile fluid is a valuable source of extracellular vesicles/exosomes that contain potentially important biomarkers. This protocol represents a robust method to isolate exosomes from human bile for further analyses including miRNA profiling.

Exosome research in the last three years has greatly extended the scope towards identification and characterization of biomarkers and their therapeutic ...

Using Multi-fluorinated Bile Acids and In Vivo Magnetic Resonance Imaging to Measure Bile Acid Transport

Along with their traditional role as detergents that facilitate fat absorption, emerging literature indicates that bile acids are potent signaling molecules that affect multiple organs; they modulate gut motility and hormone production, and alter vascular tone, glucose metabolism, lipid metabolism, and energy utilization. Changes in fecal bile acids may alter the gut microbiome and promote colon pathology including cholerrheic diarrhea and colon cancer. Key regulators of fecal bile acid composition are the small intestinal Apical Sodium-dependent Bile Acid Transporter (ASBT) and fibroblast growth factor-19 (FGF19). Reduced expression and function of ASBT decreases intestinal bile acid up-take. Moreover, in vitro data suggest that some FDA-approved drugs inhibit ASBT function. Deficient FGF19 release increases hepatic bile acid synthesis and release into the intestines to levels that overwhelm ASBT. Either ASBT dysfunction or FGF19 deficiency increases fecal bile acids and may cause chronic diarrhea and promote colon neoplasia. Regrettably, tools to measure bile acid malabsorption and the actions of drugs on bile acid transport in vivo are limited. To understand the complex actions of bile acids, techniques are required that permit simultaneous monitoring of bile acids in the gut and metabolic tissues. This led us to conceive an innovative method to measure bile acid transport in live animals using a combination of proton (1H) and fluorine (19F) magnetic resonance imaging (MRI). Novel tracers for fluorine (19F)-based live animal MRI were created and tested, both in vitro and in vivo. Strengths of this approach include the lack of exposure to ionizing radiation and translational potential for clinical research and practice.

Using Multi-fluorinated Bile Acids and In Vivo Magnetic Resonance Imaging to Measure Bile Acid Transport

Tools to diagnose bile acid malabsorption and measure bile acid transport in vivo are limited. An innovative approach in live animals is described that utilizes combined proton (1H) plus fluorine (19F) magnetic resonance imaging; this novel methodology has translational potential to screen for bile acid malabsorption in clinical practice.