This work was supported by research grants from the National Health and Medical Research Council (NHMRC) of Australia (Grant No. APP1048740 and APP1142394 to G.A.R.; APP1160323 to J.E.E.T., J.K.O., G.A.R.). Grant A. Ramm is supported by a Senior Research Fellowship from the NHMRC of Australia (Grant No. APP1061332). Manuel Fernandez-Rojo was supported by the TALENTO program of Madrid, Spain (T1-BIO-1854).

All animal experiments were performed in accordance with the Australian code for the care and use of animals for scientific purposes at QIMR Berghofer Medical Research Institute with approval from the institute animal ethics committee. Male C57BL/6 mice (15&#8722;20 weeks old) were obtained from the Animal Resources Centre, WA, Australia.
NOTE: All solutions, media, instruments, hardware, and tubes that contact the samples must be sterilized or thoroughly disinfected with a 70% ethanol solution and handled using sterile techniques to minimize the risk of culture contamination.
1. Setup of the vibratome
<ol>
	<li>Prepare sterile Krebs-Henseleit buffer solution with 2 g/L glucose (<a href="https://www.jove.com/pdf-materials/60992?refresh=1" target="_blank">Table of Materials</a>). Check that the pH of the buffer is 7.4. If the pH is higher, saturate the buffer with carbogen (95% O2 + 5% CO2) or incubate within a 5% CO2 tissue culture incubator to correct the bicarbonate buffering system. Refrigerate the sealed sterile buffer solution at 4 &#176;C.</li>
	<li>Insert the vibratome blade into the cutting arm. Make sure the blade is tightly fixed to the cutting arm, as vibrations can cause the blade to come loose.</li>
	<li>Disinfect the blade and cutting arm with a 70% ethanol solution. 
	CAUTION: Avoid contact with the blade.</li>
	<li>Disinfect the buffer tray with a 70% ethanol solution and wipe it with a sterile tissue.</li>
	<li>Insert the buffer tray into the vibratome and tighten the mounting mechanism.</li>
	<li>Set the cutting arm to the maximum available height.</li>
	<li>Set the blade angle to 10&#176; below horizontal, sloping downwards to the sample. Make sure that the blade angle is tightly fixed. 
	NOTE: The optimal blade angle may differ depending on the vibratome model and tissue characteristics.</li>
	<li>Disinfect the specimen holder with a 70% ethanol solution.</li>
	<li>Connect the cooling water for the Peltier thermoelectric cooler located under the buffer tray. 
	NOTE: Some vibratome models use an ice-bath for cooling instead.</li>
	<li>Fix the water-out tube to a drain and turn on the water. Set the cooler to 4 &#176;C. Run cooling water at a rate of &#62;400 mL/min.</li>
	<li>Fill the buffer tray almost to the top with sterile ice-cold Krebs-Henseleit buffer (with 2 g/L glucose). Leave additional Krebs-Henseleit buffer solution on ice.</li>
</ol>
2. Liver removal and preparation
<ol>
	<li>Sterilize or disinfect all surgical instruments including curved forceps, flat square-tip forceps, tweezers, surgical clamps, and scissors using autoclaving.</li>
	<li>Prior to liver removal, deeply anesthetize mice using an intraperitoneal injection of 100 mg/kg ketamine and 12.5 mg/kg xylazine. 
	NOTE: Other anesthetics may be used, or mice can be euthanized by CO2 asphyxiation/cervical dislocation if the liver is quickly removed to prevent hypoxia-induced damage.</li>
	<li>Disinfect skin surfaces on mice by wetting with a 70% ethanol solution. Secure mice on their backs with all extremities pinned to a dissecting board.</li>
	<li>Make a vertical midline incision into the skin from the base of the abdominal cavity to just above the diaphragm. 
	NOTE: Incisions towards the extremities are made to facilitate the retraction of the skin.</li>
	<li>Pull the skin back while cutting any connective tissue between the skin and abdominal cavity. Prevent hair from being transferred to the abdominal cavity as much as possible. Pin or clamp the skin away from the abdominal cavity.</li>
	<li>Using clean forceps and scissors, open the abdominal cavity and lower thoracic cavity.</li>
	<li>Removing the liver quickly without damaging the lobes
	<ol>
		<li>Using blunt tools (e.g., flat square-tip forceps), guide the upper liver lobes downwards towards the abdomen. Cut the connective tissue surrounding the upper liver lobes using scissors.</li>
		<li>Guide the liver lobes upwards towards the diaphragm. Hold the central vascular bundle of the liver with forceps. 
		NOTE: It will not affect the procedure if the central vascular bundle is damaged.</li>
		<li>Pull the central vascular bundle away from the mouse and cut the remaining connective tissue, blood vessels, etc. to remove the liver. Take care not to 1) damage the liver lobes and 2) cut into the gastrointestinal tract, as this may contaminate the liver with microorganisms.</li>
	</ol>
	</li>
	<li>Place the removed liver into a sterile 10 cm tissue culture dish half-filled with ice-cold Krebs-Henseleit buffer solution. Using a blunt instrument to guide the lobes, cut the center of the liver to divide it into separate lobes.</li>
	<li>Select one liver lobe (use the largest first), and place it flat side down onto a new sterile 10 cm dish. Keep all other liver lobes in the dish of Krebs-Henseleit buffer solution stored on ice for subsequent use.</li>
	<li>Trim around 10% of the material from the tallest edge of the liver lobe while lying flat. 
	NOTE: Trimming is important, as this tissue edge will contact the cutting blade first; therefore, a tissue edge that is relatively perpendicular to the cutting blade will prevent tissue compression that can occur because of shallow angles in the uncut liver lobes.</li>
	<li>Trim the other three tissue edges. 
	NOTE: This removes some of the fibrous Glisson&#39;s capsule and will make separation of the tissue slices easier.</li>
	<li>Mounting the liver lobe on the specimen holder
	<ol>
		<li>Place a thin layer of cyanoacrylate glue (superglue or medical/veterinary grade cyanoacrylate glue) on the specimen holder towards the front, sized slightly larger than the trimmed liver lobe. 
		NOTE: Check that the cyanoacrylate glue does not contain non-cyanoacrylate additives.</li>
		<li>Gently pick up the liver lobe with sterile forceps using a corner away from the cutting edge, then pat dry any residual buffer from the flat side of the liver lobe using sterile absorbent material.</li>
		<li>Place the large flat side of the liver lobe on the cyanoacrylate glue patch with the largest edge facing towards the front. Allow it to cure in the air for 1&#8722;2 min. 
		NOTE: Cyanoacrylate adhesives cure quickly (~2 min) in response to residual moisture and tissue proteins, and they will firmly attach the tissue to the specimen holder.</li>
	</ol>
	</li>
</ol>
3. Production of liver slices
<ol>
	<li>Place the specimen holder with the attached liver lobe into the vibratome buffer tray. Make sure the liver lobe is fully covered by buffer. 
	NOTE: If needed, top up the level of ice-cold Krebs-Henseleit buffer solution. If the buffer level obscures the cutting process, increase or decrease the level.</li>
	<li>Set the cutting speed. 
	NOTE: This may need to be empirically determined depending on the vibratome model and blade characteristics. As a starting guide, use a speed of 57 Hz/3,420 rpm. The vibratome speed can be measured using an audio spectrum analyzer application on a smartphone.</li>
	<li>Lower the cutting blade until it is located just above the liver lobe using the height dial. Run the vibrating cutting arm over the sample.</li>
	<li>Return the blade back using the crank handle in reverse. Activate the vibrations while returning the blade. After the blade has returned and is not above the sample, lower the blade height by 250 &#181;m.</li>
	<li>Repeat the cutting process until the top of the tissue is removed. Discard the first one or two slices, as these will contain Glisson&#39;s capsule and therefore do not contain much functional liver tissue compared to other slices.</li>
	<li>To cut liver slices, slowly advance the vibrating blade into the tissue by turning the handle. Use a small paintbrush (disinfected with a 70% ethanol solution) to gently guide the tissue during the cutting process.</li>
	<li>Use the paintbrush to lift the cut tissue slices and place the tissue slice into a sterile tube of ice-cold Krebs-Henseleit buffer. When not in use, keep this tube on ice between slices. 
	NOTE: Sometimes, the tissue does tear during the cutting process, but for most purposes the tissue is still usable.</li>
	<li>Repeat the cutting process until the base of the liver lobe is reached. Discard any liver slices that have visible cured glue. 
	NOTE: The cured glue is visible as white areas on the tissue slices. A typical liver lobe will only have a useable thickness of around 2 mm. Therefore, only around eight 250 &#181;m liver slices are produced from each lobe.</li>
	<li>Cut the tissue slices until near the cyanoacrylate glue. Do not cut into the glue.</li>
	<li>Repeat the whole process with other liver lobes until the required number of tissue slices is obtained.</li>
	<li>To clean the cured cyanoacrylate glue and tissue from specimen holder, either use a blade to scrape the cured glue/tissue mixture off, or soften and clean the mixture with acetone or dimethyl sulfoxide.</li>
</ol>
4. Tissue culture
NOTE: All tissue culture work must be performed in a sterile laminar flow hood.
<ol>
	<li>In a tissue culture laminar flow hood, pipette 1 mL of William&#39;s E medium containing 2.0 g/L glucose, 10% fetal bovine serum (FBS), 2 mM L-glutamine supplement, 100 U/mL penicillin, and 100 &#181;g/mL streptomycin into 12 well tissue culture plates. 
	NOTE: Other antibiotics and antifungal agents may also be added. Some studies have used additives in the tissue incubation medium (i.e., dexamethasone and insulin10), but these can interfere with cell signaling.</li>
	<li>Place the tube containing liver slices from ice into the tissue culture hood. To remove the tissue slices, gently swirl the mixture and gently tip into a sterile 10 cm dish while swirling.</li>
	<li>Using sterile sharp-point forceps and a scalpel, cut the tissue slices into roughly uniform sizes (e.g., 15 mm2). 
	NOTE: This step is performed to ensure a consistent surface area. Because mouse liver lobes are significantly different in size and some tissue slices will tear, this will produce a range of PCLS with varying surface areas. The final surface area sizes of the PCLS depend on how much material is needed in subsequent analysis applications and how many replicates are needed. Cutting large PCLS into smaller multiple pieces of the same approximate size will innately allow an increased number of experimental replicates.</li>
	<li>Using sterile forceps or a small paintbrush, transfer the cut tissue slices to the wells of a 12 well plate containing 1 mL of medium. 
	NOTE: Take care to confirm the consistency of thickness and shape of tissue slices. Sections that are darker than average suggest the tissue thickness is larger and need to be discarded. Lighter slices suggest the presence of cured cyanoacrylate glue and need to be discarded.</li>
	<li>Incubate the liver slices at 37 &#176;C and 5% CO2 under 95% humidity. 
	NOTE: Other groups have used methods to enhance oxygen delivery to PCLS, which appear to prolong tissue survival time6,10,11,12,13 (see discussion section).</li>
	<li>On the following day, change the culture medium using a manual pipette (rather than suction) to prevent the loss of tissue through suction errors. Subsequently, change the medium on the PCLS daily. The PCLS are now ready for experimental use.</li>
</ol>
5. Example application of PCLS
<ol>
	<li>At 16 h post-generation, treat PCLS with three different bile acids: glycocholic acid (GCA), taurocholic acid (TCA), and cholic acid (CA) (as sodium salts, all at a concentration of 150 &#181;M) for 2 days.</li>
	<li>After 2 days of bile acid treatment, extract mRNA by placing tissue slices in ice-cold RNA lysis buffer and quickly homogenize using beads with a bead homogenizer (<a href="https://www.jove.com/pdf-materials/60992?refresh=1" target="_blank">Table of Materials</a>).</li>
	<li>Purify the RNA using an RNA isolation kit (<a href="https://www.jove.com/pdf-materials/60992?refresh=1" target="_blank">Table of Materials</a>) and create cDNA from the purified RNA.</li>
	<li>Perform quantitative polymerase chain reaction (<a href="https://www.jove.com/pdf-materials/60992?refresh=1" target="_blank">Table of Materials</a>) using specific primers (Table 1) on the cDNA to examine gene expression.</li>
</ol>

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The authors have nothing to disclose.

The protocol demonstrates the application of murine PCLS isolation and tissue culture, and the procedures are designed to assess both viability and utility as well as examine impacts of exogenous mediators of liver pathobiology using biochemical assays, histology, and qPCR. The experimental utility of PCLS tissue culture in rodents and humans has been demonstrated in a wide range of applications, including experimental investigations in microRNA15/RNA9/protein expression<su...

The pathogenesis of most chronic liver diseases (i.e., viral hepatitis, nonalcoholic steatohepatitis, cholestatic liver injury and liver cancer) involves complex interactions between multiple different liver cell types that drive inflammation, fibrogenesis, and cancer development1,2. To understand the molecular mechanisms underlying these chronic liver-based diseases, the interactions between multiple liver cell types must be investigated. While multiple hepatic cell lines (and more recently, organoids) can be cultured in vitro, these models do not accurately emulate the complex structure, function, and cellular diversity of the hepatic microenvironment3. Furthermore, cultured liver cells (in particular, transformed cell lines) may deviate from their original source biology. Animal models are used experimentally to investigate the interactions between multiple liver cell types. However, they may become significantly reduced in scope for experimental manipulation, due to significant off-target effects in extrahepatic organs (e.g., when testing potential therapeutics).
The use of precision-cut liver slices (PCLS) in tissue culture is an experimental technique first used in drug metabolism and toxicity studies, and it involves the cutting of viable, ultrathin (around 100&#8722;250 &#181;m thick) liver slices. This permits the direct experimental manipulation of liver tissue ex vivo4. The technique bridges an experimental gap between in vivo animal studies and in vitro cell culture methods, overcoming many drawbacks of both methods (i.e., practical limits on the range of experiments that can be performed in whole animals as well as loss of structure/function and cellular diversity with in vitro cell culture methods).
Furthermore, PCLS vastly increases experimental capacity compared to whole animal studies. As one mouse can produce more than 48 liver slices, this also facilitates the use of both control and treatment groups from the same liver. In addition, the technique physically separates the liver tissue from other organ systems; therefore, it removes potential off-target effects that can occur in whole animals when testing the effects of exogenous stimuli.
In this protocol, PCLS are generated using a vibratome with a laterally vibrating blade. Other studies have successfully used a Krumdieck tissue slicer, as described in Olinga and Schuppan5. In the vibratome, lateral vibration of the blade prevents tearing of the ultrathin tissue caused by shear stress, as the blade is pushed into the tissue. Both the vibratome and Krumdieck tissue slicer work effectively without structural embedding of liver tissue, which streamlines the slicing procedure. This technique can also be used to create PCLS from diseased livers, including those from mouse models of fibrosis/cirrhosis6 and hepatic steatosis7.
In addition to demonstrating the techniques required for preparation and tissue culture of PCLS, this report also examines the viability of these ex vivo tissues by measuring adenosine triphosphate (ATP) levels and examining tissue histology to assess necrosis and fibrosis. As a representative experimental procedure, PCLS are treated with pathophysiological concentrations of three different bile acids (glycocholic, taurocholic, and cholic acids) to simulate cholestatic liver injury. In the context of cholestatic liver injury, taurocholic acid in particular has been shown to be significantly increased in both the serum and bile of children with cystic fibrosis-associated liver disease8.
Liver progenitor cells have also been treated in vitro with taurocholic acid to simulate the elevated taurocholic acid levels observed in patients, and this treatment caused increased proliferation and differentiation of liver progenitor cells towards a biliary (cholangiocyte) phenotype9. Subsequently, PCLS were treated ex vivo with elevated levels of taurocholic acid, and increased cholangiocyte markers were observed. This supports the in vitro observation that taurocholic acid drives biliary proliferation and/or differentiation in the context of pediatric cystic fibrosis-associated liver disease9.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 cm Petri Dish</td>
			<td>GREINER</td>
			<td>664160</td>
			<td>Sterile Dish</td>
		</tr>
		<tr>
			<td>12 Well Tissue Culture Plate Flat Bottom</td>
			<td>Greiner Bio-one</td>
			<td>665180</td>
			<td></td>
		</tr>
		<tr>
			<td>70% Ethanol Solution (made with AR Grade)</td>
			<td>Chem-Supply Pty Ltd</td>
			<td>EA043-20L-P</td>
			<td>Disinfection solution</td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Chem-Supply Pty Ltd</td>
			<td>AA008-2.5L</td>
			<td></td>
		</tr>
		<tr>
			<td>Cholic acid</td>
			<td>Sigma-Aldrich</td>
			<td>C1129-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Cyanoacrylate Super Glue</td>
			<td>Parfix, DuluxGroup (Australia)</td>
			<td></td>
			<td>Other brands should work</td>
		</tr>
		<tr>
			<td>Disposable Single Edge Safety Razor Blades</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection Board</td>
			<td>Made in-house</td>
			<td></td>
			<td>Sterile material over polystyrene</td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum</td>
			<td>GE Healthcare Australia Pty Ltd</td>
			<td>SH30084.02</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps sharp point 130 mm long</td>
			<td>ThermoFisher Scientific</td>
			<td>MET2115-130</td>
			<td></td>
		</tr>
		<tr>
			<td>Forma Steri-Cycle CO2 Incubator</td>
			<td>ThermoFisher Scientific</td>
			<td>371</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamine</td>
			<td>Life Technologies Australia Pty Ltd</td>
			<td>25030081</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycocholic acid hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>G2878-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>ISOLATE II RNA Mini Kit</td>
			<td>Bioline (Aust) Pty Ltd</td>
			<td>BIO-52073</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine 50 ml</td>
			<td>Provet</td>
			<td>KETAI1</td>
			<td></td>
		</tr>
		<tr>
			<td>Krebs-Henseleit Buffer with Added Glucose 2000 mg/L</td>
			<td>Sigma-Aldrich</td>
			<td>K3753</td>
			<td>Can also be made in house</td>
		</tr>
		<tr>
			<td>Laminar Flow Hood</td>
			<td></td>
			<td></td>
			<td>Hepa air filtration</td>
		</tr>
		<tr>
			<td>NanoDrop 2000/2000c Spectrophotometers</td>
			<td>ThermoFisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin, Liq 100 ml</td>
			<td>Life Technologies Australia Pty Ltd</td>
			<td>15140-122</td>
			<td></td>
		</tr>
		<tr>
			<td>Picro Sirius Red</td>
			<td>ABCAM Australia Pty Ltd</td>
			<td>ab246832</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Tips Abt 1000 &#181;l Filter Interpath</td>
			<td>Interpath</td>
			<td>24800</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Tips Abt 10 &#181;l Filter Interpath</td>
			<td>Interpath</td>
			<td>24300</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Tips Abt 200 &#181;l Filter Interpath</td>
			<td>Interpath</td>
			<td>24700</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette Tips Abt 20 &#181;l Filter Interpath</td>
			<td>Interpath</td>
			<td>24500</td>
			<td></td>
		</tr>
		<tr>
			<td>Precellys Homogeniser</td>
			<td>Bertin Instruments</td>
			<td>P000669-PR240-A</td>
			<td></td>
		</tr>
		<tr>
			<td>Protractor</td>
			<td>Generic</td>
			<td></td>
			<td>To measure blade angle</td>
		</tr>
		<tr>
			<td>Quantstudio 5 QPCR Fixed 384 Block</td>
			<td>Applied Biosystems/ ThermoFisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Blade</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel Blade Holder</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SensiFAST cDNA Synthesis Kit</td>
			<td>Bioline (Aust) PTY LTD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Small Paintbrush with Plastic Handle</td>
			<td>Mixed</td>
			<td></td>
			<td>Plastic handle resists ethanol</td>
		</tr>
		<tr>
			<td>Square-Head Foreceps</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile 50 ml Plastic Tubes</td>
			<td>Corning Falcon</td>
			<td>352098</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Clamps</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Forceps</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Pins</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scissors</td>
			<td>Mixed</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Taurochoic acid</td>
			<td>Sigma-Aldrich</td>
			<td>T-4009-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome SYS-NVSLM1 Motorized Vibroslice</td>
			<td>World Precision Instruments</td>
			<td>SYS-NVSLM1</td>
			<td>With thermoelectric cooling</td>
		</tr>
		<tr>
			<td>Williams Medium E</td>
			<td>Life Technologies Australia Pty Ltd</td>
			<td>12551032</td>
			<td>2.0 g/l glucose</td>
		</tr>
		<tr>
			<td>Xylazine 100 mg/mL 50 mL</td>
			<td>Provet</td>
			<td>XYLAZ4</td>
			<td></td>
		</tr>
	</tbody>
</table>

murine precision-cut liver slices as an ex vivo model of liver biology

Understanding the mechanisms of liver injury, hepatic fibrosis, and cirrhosis that underlie chronic liver diseases (i.e., viral hepatitis, non-alcoholic fatty liver disease, metabolic liver disease, and liver cancer) requires experimental manipulation of animal models and in vitro cell cultures. Both techniques have limitations, such as the requirement of large numbers of animals for in vivo manipulation. However, in vitro cell cultures do not reproduce the structure and function of the multicellular hepatic environment. The use of precision-cut liver slices is a technique in which uniform slices of viable mouse liver are maintained in laboratory tissue culture for experimental manipulation. This technique occupies an experimental niche that exists between animal studies and in vitro cell culture methods. The presented protocol describes a straightforward and reliable method to isolate and culture precision-cut liver slices from mice. As an application of this technique, ex vivo liver slices are treated with bile acids to simulate cholestatic liver injury and ultimately assess the mechanisms of hepatic fibrogenesis.

To determine the cell viability of PCLS over time, tissue ATP levels were measured. ATP levels are typically proportional to viability. PCLS (around 15 mm2 in area) were cultured in normal William&#39;s E medium with 10% FBS, then at specific timepoints, liver slices were removed from tissue culture and homogenized with both ATP and protein (for normalization) concentrations (<a href="https://www.jove.com/pdf-materials/60992?refresh=1" target="_blank">Table of Materials</a>) being measured (<s...

Watch this Scientific Journal Video about Murine Precision-Cut Liver Slices as an Ex Vivo Model of Liver Biology at JoVE.com

Murine Precision-Cut Liver Slices as an Ex Vivo Model of Liver Biology

This protocol provides a simple and reliable method for the production of viable precision-cut liver slices from mice. The ex vivo tissue samples can be maintained under laboratory tissue culture conditions for multiple days, providing a flexible model to examine liver pathobiology.

Understanding the mechanisms of liver injury, hepatic fibrosis, and cirrhosis that underlie chronic liver diseases (i.e., viral hepatitis, non-alcoholic ...

murine-precision-cut-liver-slices-as-an-ex-vivo-model-of-liver-biology

Research

JoVE Journal

Medicine

15.3K Views.  QIMR Berghofer Medical Research Institute. This protocol provides a simple and reliable method for the production of viable precision-cut liver slices from mice. The ex vivo tissue samples can be maintained under laboratory tissue culture conditions for multiple days, providing a flexible model to examine liver pathobiology.

Video: Murine Precision-Cut Liver Slices as an Ex Vivo Model of Liver Biology

A Human Ex Vivo Atherosclerotic Plaque Model to Study Lesion Biology

Atherosclerosis is a chronic inflammatory disease of the vasculature. There are various methods to study the inflammatory compound in atherosclerotic lesions. Mouse models are an important tool to investigate inflammatory processes in atherogenesis, but these models suffer from the phenotypic and functional differences between the murine and human immune system. In vitro cell experiments are used to specifically evaluate cell type-dependent changes caused by a substance of interest, but culture-dependent variations and the inability to analyze the influence of specific molecules in the context of the inflammatory compound in atherosclerotic lesions limit the impact of the results. In addition, measuring levels of a molecule of interest in human blood helps to further investigate its clinical relevance, but this represents systemic and not local inflammation. Therefore, we here describe a plaque culture model to study human atherosclerotic lesion biology ex vivo. In short, fresh plaques are obtained from patients undergoing endarterectomy or coronary artery bypass grafting and stored in RPMI medium on ice until usage. The specimens are cut into small pieces followed by random distribution into a 48-well plate, containing RPMI medium in addition to a substance of interest such as cytokines or chemokines alone or in combination for defined periods of time. After incubation, the plaque pieces can be shock frozen for mRNA isolation, embedded in Paraffin or OCT for immunohistochemistry staining or smashed and lysed for western blotting. Furthermore, cells may be isolated from the plaque for flow cytometry analysis. In addition, supernatants can be collected for protein measurement by ELISA. In conclusion, the presented ex vivo model opens the possibility to further study inflammatory lesional biology, which may result in identification of novel disease mechanisms and therapeutic targets.

A Human Ex Vivo Atherosclerotic Plaque Model to Study Lesion Biology

Atherosclerosis is a chronic inflammatory process. This manuscript illustrates an easy to use ex vivo model to investigate fresh carotid or coronary artery plaques. The ex vivo model allows for the investigation of potential substances on the inflammatory milieu in human atherosclerotic lesions and results can be analyzed by various methods.

Atherosclerosis is a chronic inflammatory disease of the vasculature. There are various methods to study the inflammatory compound in atherosclerotic ...

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

The use of ex vivo perfused models can mimic the physiological conditions of the liver for short periods, but to maintain normal homeostasis for an extended perfusion period is challenging. We have added the kidney to our previous ex vivo perfused liver experiment model to reproduce a more accurate physiological state for prolonged experiments without using live animals. Five intact livers and kidneys were retrieved post-mortem from sacrificed pigs on different days and perfused for a minimum of 6 hr. Hourly arterial blood gases were obtained to analyze pH, lactate, glucose and renal parameters. The primary endpoint was to investigate the effect of adding one kidney to the model on the acid base balance, glucose, and electrolyte levels. The result of this liver-kidney experiment was compared to the results of five previous liver only perfusion models. In summary, with the addition of one kidney to the ex vivo liver circuit, hyperglycemia and metabolic acidosis were improved. In addition this model reproduces the physiological and metabolic responses of the liver sufficiently accurately to obviate the need for the use of live animals. The ex vivo liver-kidney perfusion model can be used as an alternative method in organ specific studies. It provides a disconnection from numerous systemic influences and allows specific and accurate adjustments of arterial and venous pressures and flow.

Steps for the Autologous Ex vivo Perfused Porcine Liver-kidney Experiment

Study of the animal organ physiology can be achieved by performing an experimental ex vivo perfusion system. The addition of a porcine kidney, as a homeostatic organ to our previously developed ex vivo liver perfusion model can be a principal step to achieve a better physiological environment.

The use of ex vivo perfused models can mimic the physiological conditions of the liver for short periods, but to maintain normal homeostasis for an ...

Videomorphometric Analysis of Hypoxic Pulmonary Vasoconstriction of Intra-pulmonary Arteries Using Murine Precision Cut Lung Slices

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ventilation. Up to now, the underlying mechanisms are not fully understood. The major vascular segment contributing to HPV is the intra-acinar artery. This vessel section is responsible for the blood supply of an individual acinus, which is defined as the portion of lung distal to a terminal bronchiole. Intra-acinar arteries are mostly located in that part of the lung that cannot be selectively reached by a number of commonly used techniques such as measurement of the pulmonary artery pressure in isolated perfused lungs or force recordings from dissected proximal pulmonary artery segments1,2. The analysis of subpleural vessels by real-time confocal laser scanning luminescence microscopy is limited to vessels with up to 50 &#181;m in diameter3.
We provide a technique to study HPV of murine intra-pulmonary arteries in the range of 20-100 &#181;m inner diameters. It is based on the videomorphometric analysis of cross-sectioned arteries in precision cut lung slices (PCLS). This method allows the quantitative measurement of vasoreactivity of small intra-acinar arteries with inner diameter between 20-40 &#181;m which are located at gussets of alveolar septa next to alveolar ducts and of larger pre-acinar arteries with inner diameters between 40-100 &#181;m which run adjacent to bronchi and bronchioles. In contrast to real-time imaging of subpleural vessels in anesthetized and ventilated mice, videomorphometric analysis of PCLS occurs under conditions free of shear stress. In our experimental model both arterial segments exhibit a monophasic HPV when exposed to medium gassed with 1% O2 and the response fades after 30-40 min at hypoxia.

Hypoxic pulmonary vasoconstriction (HPV) is an important physiological phenomenon by which at alveolar hypoxia lung perfusion is matched to ventilation. The major vascular segment contributing to HPV is the intra-acinar artery. Here, we describe our protocol for the analysis of HPV of murine pulmonary vessels with diameters of 20-100 &mu;m.

Acute alveolar hypoxia causes pulmonary vasoconstriction (HPV) - also known as von Euler-Liljestrand mechanism - which serves to match lung perfusion to ...

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for liver transplantation die without receiving an organ transplant or are delisted for disease progression. One strategy to increase the donor pool is the utilization of marginal grafts, such as fatty livers, grafts from older donors, or donation after cardiac death (DCD). The current preservation technique of cold static storage is only poorly tolerated by marginal livers resulting in significant organ damage. In addition, cold static organ storage does not allow graft assessment or repair prior to transplantation.
These shortcomings of cold static preservation have triggered an interest in warm perfused organ preservation to reduce cold ischemic injury, assess liver grafts during preservation, and explore the opportunity to repair marginal livers prior to transplantation. The optimal pressure and flow conditions, perfusion temperature, composition of the perfusion solution and the need for an oxygen carrier has been controversial in the past.
In spite of promising results in several animal studies, the complexity and the costs have prevented a broader clinical application so far. Recently, with enhanced technology and a better understanding of liver physiology during ex vivo perfusion the outcome of warm liver perfusion has improved and consistently good results can be achieved.
This paper will provide information about liver retrieval, storage techniques, and isolated liver perfusion in pigs. We will illustrate a) the requirements to ensure sufficient oxygen supply to the organ, b) technical considerations about the perfusion machine and the perfusion solution, and c) biochemical aspects of isolated organs.

Technique of Subnormothermic Ex Vivo Liver Perfusion for the Storage, Assessment, and Repair of Marginal Liver Grafts

Marginal grafts, such as fatty livers, grafts from older donors, or livers retrieved after cardiac death (DCD) tolerate conventional, cold static storage only poorly. We developed a novel model of subnormothermic ex vivo liver perfusion for preservation, assessment, and repair of marginal liver grafts prior to transplantation.

The success of liver transplantation has resulted in a dramatic organ shortage. In most transplant regions 20-30% of patients on the waiting list for ...

The In Ovo Chick Chorioallantoic Membrane (CAM) Assay as an Efficient Xenograft Model of Hepatocellular Carcinoma

The chick chorioallantoic membrane (CAM) begins to develop by day 7 after fertilization and matures by day 12. The CAM is naturally immunodeficient and highly vascularized, making it an ideal system for tumor implantation. Furthermore, the CAM contains extracellular matrix proteins such as fibronectin, laminin, collagen, integrin alpha(v)beta3, and MMP-2, making it an attractive model to study tumor invasion and metastasis. Scientists have long taken advantage of the physiology of the CAM by using it as a model of angiogenesis. More recently, the CAM assay has been modified to work as an in vivo xenograft model system for various cancers that bridges the gap between basic in vitro work and more complex animal cancer models. The CAM assay allows for the study of tumor growth, anti-tumor therapies, and pro-tumor molecular pathways in a biologically relevant system that is both cost- and time-effective. Here, we describe the development of CAM xenograft model of hepatocellular carcinoma (HCC) with embryonic survival rates of up to 93% and reliable tumor take leading to growth of three-dimensional, vascularized tumors.

The In Ovo Chick Chorioallantoic Membrane CAM Assay as an Efficient Xenograft Model of Hepatocellular Carcinoma

The chick chorioallantoic membrane (CAM) is immunodeficient and highly vascularized, making it a natural in vivo model of tumor growth and angiogenesis. In this protocol, we describe a reliable method of growing three-dimensional, vascularized hepatocellular carcinoma (HCC) tumors using the CAM assay.

The chick chorioallantoic membrane (CAM) begins to develop by day 7 after fertilization and matures by day 12. The CAM is naturally immunodeficient and ...

Assessment of Plasma Coagulation on Liver Tissue in a Large Animal Model In Vivo

Plasma coagulation as a form of electrocautery is used in liver surgery for decades to seal the large liver cut surface after major hepatectomy to prevent hemorrhages at a later stage. The exact effects of plasma coagulation on liver tissue are only poorly examined. In our porcine model, the coagulation effects can be examined close to the clinical application. A combined laser Doppler flowmeter and spectrophotometer documents microcirculation changes during coagulation at 8 mm tissue depth noninvasively, providing quantifiable information about hemostasis beyond the subjective clinical impression. The temperature at coagulation site is assessed with an infrared thermometer prior and post coagulation and with a thermographic camera during coagulation, a measurement of the gas beam temperature is not possible due to the upper threshold of the devices. The depth of coagulation is measured microscopically on hematoxylin/eosin stained sections after calibration with an object micrometer and gives an exact information about the power setting-coagulation depth-relation. The sealing effect is examined on the bile ducts as it is not possible for a plasma coagulator to seal larger blood vessels. Burst pressure experiments are carried out on explanted organs to rule out blood pressure related effects.

Assessment of Plasma Coagulation on Liver Tissue in a Large Animal Model In Vivo

Here we present a protocol to experimentally assess plasma coagulation in liver tissue in vivo. In a porcine model, microcirculation is examined by laser Doppler, coagulation depth is measured histologically, temperature at coagulation site by infrared thermometer and thermographic camera, and duct sealing effect is documented by burst pressure experiments.

Plasma coagulation as a form of electrocautery is used in liver surgery for decades to seal the large liver cut surface after major hepatectomy to prevent ...

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, normothermic ex vivo liver perfusion (NEVLP) has been introduced as a method to evaluate and modify organ function. NEVLP has many advantages in comparison to hypothermic and subnormothermic perfusion including reduced preservation injury, restoration of normal organ function under physiologic conditions, assessment of organ performance, and as a platform for organ repair, remodeling, and modification. Both murine and porcine NEVLP models have been described. We demonstrate a rat model of NEVLP and use this model to show one of its important applications &#8212; the use of a therapeutic molecule added to liver perfusate. Catalase is an endogenous reactive oxygen species (ROS) scavenger and has been demonstrated to decrease ischemia-reperfusion in the eye, brain, and lung. Pegylation has been shown to target catalase to the endothelium. Here, we added pegylated-catalase (PEG-CAT) to the base perfusate and demonstrated its ability to mitigate liver preservation injury. An advantage of our rodent NEVLP model is that it is inexpensive in comparison to larger animal models. A limitation of this study is that it does not currently include post-perfusion liver transplantation. Therefore, prediction of the function of the organ post-transplantation cannot be made with certainty. However, the rat liver transplant model is well established and certainly could be used in conjunction with this model. In conclusion, we have demonstrated an inexpensive, simple, easily replicable NEVLP model using rats. Applications of this model can include testing novel perfusates and perfusate additives, testing software designed for organ evaluation, and experiments designed to repair organs.

A Small Animal Model of Ex Vivo Normothermic Liver Perfusion

There is a significant liver donor shortage, and criteria for liver donors have been expanded. Normothermic ex vivo liver perfusion (NEVLP) has been developed to evaluate and modify organ function. This study demonstrates a rat model of NEVLP and tests the ability of pegylated-catalase, to mitigate liver preservation injury.

There is a significant shortage of liver allografts available for transplantation, and in response the donor criteria have been expanded. As a result, ...

An Ex Vivo Tissue Culture Model for Fibrovascular Complications in Proliferative Diabetic Retinopathy

Diabetic retinopathy (DR) is the most common microvascular complication of diabetes and one of the leading causes of blindness in working-age adults. No current animal models of diabetes and oxygen-induced retinopathy develop the full-range progressive changes manifested in human proliferative diabetic retinopathy (PDR). Therefore, understanding of the disease pathogenesis and pathophysiology has relied largely on the use of histological sections and vitreous samples in approaches that only provide steady-state information on the involved pathogenic factors. Increasing evidence indicates that dynamic cell-cell and cell-extracellular matrix (ECM) interactions in the context of three-dimensional (3D) microenvironments are essential for the mechanistic and functional studies towards the development of new treatment strategies. Therefore, we hypothesized that the pathological fibrovascular tissue surgically excised from eyes with PDR could be utilized to reliably unravel the cellular and molecular mechanisms of this devastating disease and to test the potential for novel clinical interventions. Towards this end, we developed a novel method for 3D ex vivo culture of surgically-excised patient-derived fibrovascular tissue (FT), which will serve as a relevant model of human PDR pathophysiology. The FTs are dissected into explants and embedded in fibrin matrix for ex vivo culture and 3D characterization. Whole-mount immunofluorescence of the native FTs and end-point cultures allows thorough investigation of tissue composition and multicellular processes, highlighting the importance of 3D tissue-level characterization for uncovering relevant features of PDR pathophysiology. This model will allow the simultaneous assessment of molecular mechanisms, cellular/tissue processes and treatment responses in the complex context of dynamic biochemical and physical interactions within the PDR tissue architecture and microenvironment. Since this model recapitulates PDR pathophysiology, it will also be amenable for testing or developing new treatments.

Here, we present a protocol to study the pathophysiology of proliferative diabetic retinopathy by using patient-derived, surgically-excised, fibrovascular tissues for three-dimensional native tissue characterization and ex vivo culture. This ex vivo culture model is also amenable for testing or developing new treatments.

Diabetic retinopathy (DR) is the most common microvascular complication of diabetes and one of the leading causes of blindness in working-age adults. No ...

Vinyl Chloride and High-Fat Diet as a Model of Environment and Obesity Interaction

Vinyl chloride (VC), an abundant environmental contaminant, causes steatohepatitis at high levels, but is considered safe at lower levels. Although several studies have investigated the role of VC as a direct hepatotoxicant, the concept that VC modifies sensitivity of the liver to other factors, such as nonalcoholic fatty liver disease (NAFLD) caused by high-fat diet (HFD) is novel. This protocol describes an exposure paradigm to evaluate the effects of chronic, low-level exposure to VC. Mice are acclimated to low-fat or high-fat diet one week prior to the beginning of the inhalation exposure and remain on these diets throughout the experiment. Mice are exposed to VC (sub-OSHA level: &#60;1 ppm) or room air in inhalation chambers for 6 hours/day, 5 days/week, for up to 12 weeks. Animals are monitored weekly for body weight gain and food consumption. This model of VC exposure causes no overt liver injury with VC inhalation alone. However, the combination of VC and HFD significantly enhances liver disease. A technical advantage of this co-exposure model is the whole-body exposure, without restraint. Moreover, the conditions more closely resemble a very common human situation of a combined exposure to VC with underlying nonalcoholic fatty liver disease and therefore support the novel hypothesis that VC is an environmental risk factor for the development of liver damage as a complication of obesity (i.e., NAFLD). This work challenges the paradigm that the current exposure limits of VC (occupational and environmental) are safe. The use of this model can shed new light and concern on the risks of VC exposure. This model of toxicant-induced liver injury can be used for other volatile organic compounds and to study other interactions that may impact the liver and other organ systems.

The goal of this protocol was to develop a murine model of low-level toxicant exposure that does not cause overt liver injury but rather exacerbates pre-existing liver damage. This paradigm better recapitulates human exposure and the subtle changes that occur upon exposure to toxicant concentrations that are considered safe.

Vinyl chloride (VC), an abundant environmental contaminant, causes steatohepatitis at high levels, but is considered safe at lower levels. Although ...

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

The visualization of murine lung tissue provides valuable structural and cellular information regarding the underlying airway and vasculature. However, the preservation of pulmonary vessels that truly represents physiological conditions still presents challenges. In addition, the delicate configuration of murine lungs result in technical challenges preparing samples for high-quality images that preserve both cellular composition and architecture. Similarly, cellular contractility assays can be performed to study the potential of cells to respond to vasoconstrictors in vitro, but these assays do not reproduce the complex environment of the intact lung. In contrast to these technical issues, the precision-cut lung slice (PCLS) method can be applied as an efficient alternative to visualize lung tissue in three dimensions without regional bias and serve as a live surrogate contractility model for up to 10 days. Tissue prepared using PCLS has preserved structure and spatial orientation, making it ideal to study disease processes ex vivo. The location of endogenous tdTomato-labeled cells in PCLS harvested from an inducible tdTomato reporter murine model can be successfully visualized by confocal microscopy. After exposure to vasoconstrictors, PCLS demonstrates the preservation of both vessel contractility and lung structure, which can be captured by a time-lapse module. In combination with the other procedures, such as western blot and RNA analysis, PCLS can contribute to the comprehensive understanding of signaling cascades that underlie a wide variety of disorders and lead to a better understanding of the pathophysiology in pulmonary vascular diseases.

Precision Cut Lung Slices as an Efficient Tool for Ex vivo Pulmonary Vessel Structure and Contractility Studies

Presented here is a protocol for preserving the vascular contractility of PCLS murine lung tissue, resulting in a sophisticated three-dimensional image of the pulmonary vasculature and airway, which can be preserved for up to 10 days that is susceptible to numerous procedures.

The visualization of murine lung tissue provides valuable structural and cellular information regarding the underlying airway and vasculature. However, ...