We thank everyone assisting in this project, especially Karolina Guja and Dr. Daniela Liberati, for maintaining the organoids and guaranteeing reproducible quality over time. We thank Eva Breuer and Anurag Gupta from the Hepatobiliary and Transplant Surgery Lab of the University Hospital of Z&#252;rich for their support concerning anesthesia. This work was supported by the Surgical Department program &#34;Personenf&#246;rderung&#34; to G.F.H and F.H.; the St. Clara Forschung AG and the &#34;Stiftung zur Krebsbek&#228;mpfung&#34; to G.F.H. and University of Basel, Research Fund Junior Researcher (3MS1087) to M.C-LL.

Liver samples used for organoid generation were obtained from patients operated at the University Hospital of Basel (USB) following written informed consent under approval by the Ethical Committee of Northwestern and Central Switzerland (BASEC number 2019-02118). All mouse experiments were approved by and performed in accordance with the guidelines and regulations of the Animal Care Committee of Kanton Basel-Stadt, Switzerland (3123-33896). All mice were of the non-obese diabetic (NOD) SCID gamma mouse strain and therefore genetically modified to be immunosuppressed; only male animals were used. See Table 1 and Table of Materials for details related to solutions, reagents, materials, and instruments used in the protocol. See Figure 1 for an overview of the experiment.
1. Mouse husbandry
<ol>
	<li>Breed and maintain mice in the animal facility under specific pathogen-free conditions on a 12 h day and 12 h night schedule with ad libitum access to food and drinking water. 
	NOTE: Implantation of the organoids is performed at an age of 6-8 weeks.</li>
</ol>
2.&#160;Patient and clinical samples
<ol>
	<li>Place 12-, 24-, and 6-well plates in the incubator at 37 &#176;C before starting. If possible, do this 1 day in advance.
	<ol>
		<li>Collect the tissue within 1 h after resection in RPMI 1640 medium and transport it to the laboratory to start the dissociation process immediately. Place the tissue in a 10 cm Petri dish (prewarmed dish) and add a sufficient amount of sterile saline to avoid the tissue drying out.</li>
	</ol>
	</li>
	<li>Liver cancer tissue dissociation for the generation of organoids
	<ol>
		<li>Mince the tissue into small fragments of 1-2 mm2 using scalpels. Use additional tissue forceps to stabilize the tissue in case cutting is difficult.</li>
		<li>Place the tissue pieces in a 15 mL tube with 5 mL of dissociation buffer (physiological pH). Place at 37 &#176;C (warming cabinet/incubator) in a rotation device.</li>
		<li>Mix by pipetting every 10 min and leave for approximately 1-1.5 h, repeating the mixing process every 10 min. 
		NOTE: Mixing may require cutting the end of the pipette to increase the orifice diameter, thereby preventing oversize tissue from getting stuck in the pipette tip.</li>
		<li>Collect 10 &#181;L of cell-dissociation buffer mixture from step 2.2.3 in a 0.5 mL plastic tube, add 10 &#181;L of Trypan Blue, and use the automated cell counter to assess cell viability and cell clusters. If clusters are visible, repeat the mixing process (step 3.3.); otherwise, proceed with the next step.</li>
		<li>Take a 50 mL tube, place a 100 &#181;m cell strainer on the top, and pass the mix through it. With the plunger of a 5 mL syringe, smash all remaining tissue fragments through the filter. Wash everything with 5 mL of Ad-DF+++ and be sure to collect the drop from the bottom of the filter.</li>
		<li>Transfer the filtrate to a 15 mL conical tube. Centrifuge for 15 min at 300 &#215; g.</li>
		<li>Aspirate the supernatant carefully, taking care to not touch the pellet. Add 5 mL of Red Blood cell lysis buffer. Incubate for 5-10 min at room temperature.</li>
		<li>Centrifuge for 5 min at 300 &#215; g. Remove the supernatant and resuspend in 1 mL of advanced medium.</li>
	</ol>
	</li>
	<li>Counting cells and assessing cell viability
	<ol>
		<li>Add 10 &#181;L of Trypan blue to a 1.5 mL tube and mix with 10 &#181;L of the cell suspension. Place 10 &#181;L of the mix on the counting slides and count the cells.</li>
		<li>Separate the necessary number of cells for the organoid generation and centrifuge for 5 min at 300 &#215; g. Generally, seed 80,000 cells per dome and 20 &#181;L of basement membrane matrix per dome (10 drops per well per 6-well plate).</li>
		<li>If any excess cells are remaining in suspension, suspend the pellet in Cell Suspension Freezing medium (10% DMSO in FBS), transfer it into a cryotube, and place it in a freezing container. Immediately transfer the tube to a -80 &#176;C freezer overnight. The next day, transfer into liquid nitrogen until needed.</li>
	</ol>
	</li>
	<li>Plating for organoid generation
	<ol>
		<li>Remove the supernatant. Place the tube on ice and resuspend the pellet with a basement membrane matrix.</li>
		<li>Take the prewarmed plate out of the incubator (prewarmed overnight). Make small drops in the plate (20 &#181;L of basement membrane matrix per dome [10 drops per well]). Invert the plate and leave the plate for 5 min in the flow cabinet.</li>
		<li>After 5 min, transfer the plate to the 37 &#176;C incubator and leave the plate for ~20 min, allowing the matrix to solidify. Add 2 mL of organoid culture medium6,7.</li>
	</ol>
	</li>
	<li>Maintenance of organoid culture
	<ol>
		<li>Replace the medium every 2 days. Once the organoids are confluent in the basement membrane matrix dome and ready for expansion, calculate the splitting ratio e.g., 2x or 3x accordingly. 
		NOTE: Record this information; it will be needed in the next steps.</li>
		<li>Remove the medium from the corner of the well using a P1000 tip, taking care not to touch the matrix domes.</li>
		<li>Add 100 &#181;L of 0.25% trypsin-EDTA per dome, scrape it, collect it into a 15 mL tube, and add again a small amount of trypsin into the well to collect everything. Pipette it ~10x.</li>
		<li>Incubate for 3 min in the 37 &#176;C water bath and pipette 10x to dissociate the cells with a P1000 tip. Repeat this step every 3 min for a maximum of 10 min.</li>
		<li>Add at least the same volume of Dulbecco&#39;s modified Eagle&#39;s medium (DMEM)/10% Fetal bovine serum (FBS)/1% Penicillin-Streptomycin (Pen-Strep) medium and mix up and down slowly 2x. Fill up with DPBS.</li>
		<li>Centrifuge at 300 &#215; g for 5 min and carefully remove the supernatant. Resuspend according to the splitting ratio decided based on the organoid confluency e.g., 2x or 3x the initial amount of basement membrane matrix.</li>
		<li>Take the prewarmed plate out of the incubator (prewarmed overnight). Make small drops in the plate (20 &#181;L of basement membrane matrix per dome [10 drops per well]). Invert the plate and leave the plate for 5 min in the flow cabinet.</li>
		<li>After 5 min, transfer the plate to the 37 &#176;C incubator. Leave the plate for ~20 min, allowing the matrix to solidify. Add 2 mL of organoid culture medium.</li>
	</ol>
	</li>
	<li>Preparation of organoids for making a formalin-fixed paraffin-embedded (FFPE) block
	<ol>
		<li>Remove the medium from the corner of the well using a P1000 tip, taking care not to touch the matrix domes. Add 200 &#181;L of Dispase (2 mg/mL) and disrupt the drops with a P1000 pipette tip. Leave in the incubator for 80 min.</li>
		<li>Add 2 mL of Dulbecco&#39;s phosphate-buffered saline (DPBS) to the well, transfer it to a 15 mL conical tube, and fill it up with DPBS. Centrifuge at 300 &#215; g for 5 min, aspirate the supernatant, and resuspend the pellet in 1 mL of 4% paraformaldehyde. Refrigerate for at least 1 h.</li>
		<li>Centrifuge at 300 &#215; g for 5 min, aspirate the paraformaldehyde, and resuspend the pellet with the specimen-processing gel to create a drop. Wait for 40 min for the drops to solidify. Place the gel drop containing the cells in a histology cassette. Move the cassette to dehydrate and afterward embed it in paraffin for cutting and H/E staining8,9.</li>
	</ol>
	</li>
	<li>Preparation of organoids for injection
	<ol>
		<li>Take the plate containing the growing organoid out of the incubator and place it in the flow cabinet.</li>
		<li>Follow steps 2.5.2 to 2.5.4.</li>
		<li>Add 2 mL of DMEM and mix.</li>
		<li>Add 10 &#181;L of Trypan blue to a 1.5 mL tube and mix with 10 &#181;L of cell suspension.</li>
		<li>Place 10 &#181;L of the mix on the counting slides and count the cells. Calculate the average cell count of both chambers. If the difference is too big between the two chambers, consider redoing the count.</li>
		<li>Calculate the volume of cell suspension required for 106 cells and dispense this volume to 1.5 mL tubes. Centrifuge at 300 &#215; g for 5 min and place the tubes on ice.</li>
		<li>Carefully remove the supernatant and add 5 &#181;L of DMEM and 5 &#181;L of basement membrane matrix. Carefully mix the suspension by pipetting while keeping the suspension cold during the process.</li>
		<li>Place the 1.5 mL tubes containing the organoids on ice during the transport to the mice and proceed with injection.</li>
	</ol>
	</li>
</ol>
3. Injection of organoid into mouse
<ol>
	<li>Perioperative steps
	<ol>
		<li>Remove the experimental animal from the main cage and place it into an isolated, fresh cage with water.</li>
		<li>Anesthetize the mouse with 1.5% to 2.5% Isoflurane.</li>
		<li>Perform a subcutaneous injection into the neck with 10 &#181;L of Buprenorphine (0.3 mg/mL) 30 min prior to the start of the surgery.</li>
		<li>Immediately prior to the start of the surgery, carefully paint the eyes of animals with Carbomerum (e.g., Lacrinorm) to avoid any dryness of the eyes during surgery. Do not touch the cornea during the application to avoid any erosion of the cornea.</li>
		<li>Place the animals onto a clean and sterile surface and handle them with fresh plastic gloves, hairnet, and a special laboratory working coat used only in the mouse facility.</li>
		<li>Use two heat-emitting headlights close to the animal to provide a sufficient view. If there is any doubt or concern about hypothermia during the surgery, use another heating lamp or heating mat.</li>
	</ol>
	</li>
	<li>Laparotomy 
	NOTE: Before the surgical procedure started, the depth of the anesthesia was checked by the toe pinch test. During the procedure, adequate anesthesia was provided through routine checks and monitoring of the respiration rate and adjusted if there was a relevant increase. The surgical equipment was cleaned and disinfected according to the in-house guidelines of our animal facility.
	<ol>
		<li>Place the anesthetized mouse in the supine position, extend the four extremities, and fix them with tape. Remove the fur by applying a depilatory cream and use a wet tissue to remove the excess cream and fur. 
		NOTE: Depilation works best if a cotton swab is used to massage the cream into the fur against the grain.</li>
		<li>Localize the linea alba, which is the target for the laparotomy. Identify the superior and inferior epigastric arteries as well. Take care not to damage the vessels to avoid injuries.</li>
		<li>Make an incision along the linea alba with forceps and microsurgical scissors from the lower abdomen to the xiphoid cartilage.</li>
		<li>Insert the retractors and adjust the direction of traction for optimal exposure of the upper abdomen (Figure 2A,B). 
		NOTE: We use bent paper clips as retractors with attached rubber bands for fixation.</li>
	</ol>
	</li>
	<li>Injection of organoids
	<ol>
		<li>Place a clean napkin right next to the incision and put some drops of saline onto it. Exteriorize the small intestine and the cecum and place it on the wet napkin.</li>
		<li>Expose the right superior lobe (RSL) until an injection is possible.</li>
		<li>Prepare the Hamilton syringe, rinse it with water, and place a mark 2-3 mm from the tip as a reference for the injection.</li>
		<li>Draw up the syringe with the prepared organoid mixture and inject the needle until the mark. Slowly inject the organoids in a steady position; do not move the needle during this time.</li>
		<li>Wait for 15-20 s to avoid backflow of the mixture. Remove the needle and put some pressure with a cotton swab on the injection site. Check the abdomen for a rupture of the liver lobe and backflow. Rinse the abdomen with 0.5 mL of sterile saline and close the abdomen with a running suture using a resorbable suture. 
		NOTE: The surgical site was covered by a sterile drape. The surgical instruments used during the procedure were primarily cleaned with water and cleaning agents to remove macroscopic remnants. The instruments were then processed in a sterilization solution. Consumables like threads, gloves or cotton swabs were only used once for hygienic reasons and were sterile.</li>
	</ol>
	</li>
</ol>
4. Minor resection (30% of liver tissue)
<ol>
	<li>Perform the laparotomy as stated above in step 3.2.</li>
	<li>Use moist cotton swabs and push the ML cranially to create more space in the surgical field of the LLL.</li>
	<li>Flip the LLL cranially and expose the caudate lobe (CL) and ligament, which is habitually present between the CL and LLL (Figure 2C).</li>
	<li>Cut the ligament completely and insert the ligature (non-resorbable thin thread, e.g. Prolene 5-0) below the LLL. To ensure that it stays in this position, moisten the ligature and move it towards the right upper quadrant.</li>
	<li>Flip the LLL caudally. Guide the left free end of the ligature around the left tip of the LLL. Guide the right free end of the ligature around the right tip of the LLL. Apply caudal traction on the LLL with a cotton swab and ensure the correct and central positioning of the ligature.</li>
	<li>Tie a double half knot. Secure the knot with a second half knot tied in the other direction without fully tightening it. Tighten the knots a little bit, check the correct placement of the ligature, and reposition it to guarantee good positioning.</li>
	<li>Tighten the knot completely by holding the cranial free end in position with blunt-tipped forceps and pull the caudal free end with another pair of forceps (Figure 2D). Secure the knot with a second half knot tied in the opposite direction.</li>
	<li>Resect the liver lobe. Cut close to the ligature without cutting it.</li>
</ol>
5. Major resection (65% of liver tissue)
<ol>
	<li>Perform the minor resection of the LLL as stated above (steps 3.4.1-3.4.8).</li>
	<li>Transect the falciform ligament to mobilize the ML. Flip the lobe cranially. Insert the ligature below the lobe and flip the lobe caudally.</li>
	<li>Loop the free ends around the lobule. Repeat placement of the ligature in case of misplacement during manipulation. Once the ligature is positioned correctly, place a double half knot, tie it slowly, and reposition the ligature if needed.</li>
	<li>Tie a second half knot in the opposite direction and tighten it completely. Add a third half knot in the opposite direction.</li>
	<li>Resect the lobe.</li>
	<li>Flush the abdomen with sterile saline solution and place the intestines in their anatomical place.</li>
</ol>
6. Closure of the abdomen
<ol>
	<li>Grab the skin and peritoneum with forceps and make a stitch with a resorbable suture (e.g., Monocryl 5-0) on the top end of the incision line. Be sure to exit on the peritoneal side.</li>
	<li>Perform the second stitch on the counterpart of the incision, exiting on the skin side. Tie the knot tightly while leaving the free end without the needle short and the end with the needle long. Cut the short end to a length of 2 mm.</li>
	<li>Continue the running suture and finish with a knot at the bottom end of the incision.</li>
	<li>Clean the abdomen of the mouse of any remaining blood.</li>
</ol>
7. Postoperative care
<ol>
	<li>After the closure of the abdomen, inject another 10 &#181;L of Buprenorphine (0.3 mg/mL) subcutaneously into the neck. Place the animals in the previously used, isolated cage, partially placed on a heating mat. Provide some easily accessible food on the floor of the cage. 
	NOTE: Do not leave the animal unattended until it has regained consciousness.</li>
	<li>Carry out postoperative controls at 2 h and 6 h after the surgery. Closely observe the behavior with special attention to any sign of pain; administer another 10 &#181;L of Bupivacaine if needed.</li>
	<li>Once the mouse is fully recovered, but not prior to 6 h after surgery and not later than 24 h after surgery, re-introduce it to the main cage. Closely monitor the behavior of all the involved animals regarding any aggression or other suspicious behavior. When in doubt of any postoperative weakness of the animal, leave it isolated overnight to recover until final re-introduction.</li>
	<li>After the final postoperative control 1 day after the surgery, proceed with the regular weighing of the animals. 
	NOTE: Analgesia was administered 4 and 8 hours postoperatively. Further monitoring was performed at 12, 24 and 48 hours postoperatively, and mice were checked for any kind of pain or distress. If any signs of pain were observed, another dose of Buprenorphine was administered.</li>
</ol>
8. Monitoring tumor growth
<ol>
	<li>Immobilize the mice using isoflurane anesthesia and use sonography to visualize tumor implantation beginning 2 weeks after injection10.</li>
	<li>Euthanize the mice using CO2.</li>
</ol>

Patient&#45;Derived Hepatocarcinoma Cell Organoids for Liver Regeneration Study

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L., 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=Development+of+a+PD-L1-expressing+orthotopic+liver+cancer+model:+implications+for+immunotherapy+for+hepatocellular+carcinoma.">Development of a PD-L1-expressing orthotopic liver cancer model: implications for immunotherapy for hepatocellular carcinoma.</a> Liver Cancer. 8 (3), 155-171 (2019).</li><li>Nuciforo, S., 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=Organoid+models+of+human+liver+cancers+derived+from+tumor+needle+biopsies.">Organoid models of human liver cancers derived from tumor needle biopsies.</a> Cell Reports. 24 (5), 1363-1376 (2018).</li><li>Broutier, L., 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=Human+primary+liver+cancer-derived+organoid+cultures+for+disease+modeling+and+drug+screening.">Human primary liver cancer-derived organoid cultures for disease modeling and drug screening.</a> Nature Medicine. 23 (12), 1424-1435 (2017).</li><li>Qin, C., 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+cutting+and+floating+method+for+paraffin-embedded+tissue+for+sectioning.">The cutting and floating method for paraffin-embedded tissue for sectioning.</a> Journal of Visualized Experiments: JoVE. (139), (2018).</li><li>Feldman, A. T., Wolfe, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+processing+and+hematoxylin+and+eosin+staining.">Tissue processing and hematoxylin and eosin staining.</a> Methods in Molecular Biology. 1180, 31-43 (2014).</li><li>Bou About, 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=Ultrasound-guided+approaches+to+improve+orthotopic+mouse+xenograft+models+for+hepatocellular+carcinoma.">Ultrasound-guided approaches to improve orthotopic mouse xenograft models for hepatocellular carcinoma.</a> Current Protocols in Mouse Biology. 9 (2), 62 (2019).</li><li>Janssen, B. J. A., Smits, J. F. 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All authors have no financial or personal relationships with other people or organizations that could inappropriately influence their work. This includes any potential conflicts of interest, such as financial interests, affiliations, or relationships that might affect their ability to present or interpret data objectively.

Critical steps in the protocol 
Mouse anesthesia 
The goal of the anesthesia is to convert the mouse safely into a state in which it will tolerate surgical manipulation. At the same time, the effect of the anesthesia should not be too strong, to prevent cardiopulmonary arrest and ensuing death. Inhalant anesthesia is easier to dose, allowing the researcher to adjust the dose by manipulating the flow rate of the vaporizer. The effect of the change is quick. This helps prevent a ...

Recurrence following hepatocellular carcinoma (HCC) treatment is a significant challenge, impacting over 70% of patients undergoing surgical resection1,2,3. This recurrence may result from undetected micro-metastasis (multicentric tumor) or the development of de novo cancer4. Both clinical and experimental investigations propose that the liver regeneration process triggered by surgical resection could activate latent micro-metastases, contributing to tumor recurrence.
Further research is necessary to compare the gene and protein expression in normal and malignant liver cells within a regenerative liver environment. In addition to elucidating the mechanisms underlying recurrence, identifying targetable signaling pathways specific to cancer cells holds the potential for substantial therapeutic advancements. This approach aims to reconcile the seemingly contradictory notions of an anti-tumor microenvironment and favorable conditions for liver regeneration.
Previous models have used orthotopic injection of HCC cell lines to investigate the impact of liver regeneration5. This pioneering model employs patient-derived HCC organoids, offering several advantages over traditional cell lines. Organoids maintain diverse cell populations, mirroring the model template&#39;s structure and function, unlike cell lines and spheroids. Their genetic diversity enhances representativeness. Additionally, organoids provide stability for extended culture, enabling prolonged intervention studies. We recommend RSL injection due to its superior regeneration potential compared to the right inferior lobe (RIL). The method described here provides a platform using cancer organoids with pronounced advantages compared to cancer cell lines to investigate the aforementioned research question.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Advanced DMEM/F12 (adDMEM/F12)</td>
			<td>Life Technologies</td>
			<td>12634-034</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A3733-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>Bupaq Buprenorphinum 0.3 mg pro 1 mL</td>
			<td>Streuli Tiergesundheit AG</td>
			<td>1121915AB</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5810R</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase IV</td>
			<td>Worthington</td>
			<td>LS004189</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Matrigel Growth Factor Reduced (GFR) Basement Membrane Matrix (10 mL)</td>
			<td>Corning (Merk)</td>
			<td>CLS356231-1EA</td>
			<td></td>
		</tr>
		<tr>
			<td>Countess II Automated Cell Counter</td>
			<td>Thermo Fisher</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I Type IV from Bovine (DNAse)</td>
			<td>Sigma-Aldrich</td>
			<td>D5025-150KU</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM (1x)</td>
			<td>Gibco</td>
			<td>41965-039</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS, no calcium, no magnesium (500 mL)</td>
			<td>Gibco</td>
			<td>14190-094</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS) (South America). Heat Inactivated &#8211; (500 mL)</td>
			<td>Lubio Science</td>
			<td>S181H-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Glutamax (100x)</td>
			<td>Gibco</td>
			<td>9149793</td>
			<td></td>
		</tr>
		<tr>
			<td>Grant SUB Aqua Pro Water Bath</td>
			<td>Grant Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES (1 M)</td>
			<td>Thermo Fisher</td>
			<td>15630056</td>
			<td></td>
		</tr>
		<tr>
			<td>Histogel</td>
			<td>Thermo Fisher</td>
			<td>R904012</td>
			<td>specimen-processing&#160; gel</td>
		</tr>
		<tr>
			<td>Hyaluronidase Type IV from sheep (Tested)</td>
			<td>Sigma-Aldrich</td>
			<td>H6254-500MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted Microscope Olympus CKX53</td>
			<td>Olympus</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MacsMix Tube Rotator</td>
			<td>Miltenyi Biotec</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin-Glutamine (100x)</td>
			<td>Gibco</td>
			<td>10378-016</td>
			<td></td>
		</tr>
		<tr>
			<td>Red Blood Cell Lysis</td>
			<td>Roche</td>
			<td>11814389001</td>
			<td></td>
		</tr>
		<tr>
			<td>RPMI 1640 Medium</td>
			<td>Gibco</td>
			<td>72400-021</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA (0.25%), phenol red (100 mL)</td>
			<td>Gibco</td>
			<td>25200-056</td>
			<td></td>
		</tr>
		<tr>
			<td>Vitaris CO2 Incubator</td>
			<td>Vitaris AG</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Y-27632 dihydrochloride (Rock Inhibitor)</td>
			<td>Abmole Bioscience</td>
			<td>M1817</td>
			<td></td>
		</tr>
	</tbody>
</table>

a hepatocellular cancer patient&#45;derived organoid xenograft model to investigate impact of liver regeneration on tumor growth

Recurrence poses a notable challenge after hepatocellular carcinoma (HCC) treatment, impacting more than 70% of patients who undergo surgical resection. Recurrence stems from undetected micro-metastasis or de novo cancer, potentially triggered by postsurgical liver regeneration. Prior research employed HCC cell lines in orthotopic models to study the impact of liver regeneration, but their limited validity prompted the need for a more representative model. Here, we introduce a novel approach utilizing patient-derived HCC organoids to investigate the influence of liver regeneration on HCC.
Patient tumor tissues are processed to create tumor organoids, embedded in a three-dimensional basement membrane matrix, and cultured in a liver-specific medium. One million organoids are injected into the right superior lobe (RSL) of immunodeficient mice, confirming macroscopic tumor growth through sonography. The intervention group undergoes resection of the left lateral lobe (LLL) (30% of total liver volume) or additionally, the middle lobe (ML) (65% of total liver volume) to induce liver regeneration within the tumor site. The control group experiences re-laparotomy without liver tissue resection. After 2 weeks, both groups undergo tumor and normal tissue explantation.
In conclusion, this patient-derived HCC organoid model offers a robust platform to investigate the impact of liver regeneration post-cancer resection. Its multi-cellular composition, genetic diversity, and prolonged culture capabilities make it an invaluable tool for studying HCC recurrence mechanisms and potential interventions.

We looked at the contribution of the single liver lobes to the total liver volume and found that the LLL represents 33% of the total liver volume. The ML represents 32% of the total liver volume, the RSL 13%, the RIL 10%, and the CL 10% (Table 2). The boxplots show that the relative contribution of the liver lobes is comparable between the different mice of the same strain (Figure 3B). Hence, a resection of the LLL results in a resection of 35% of the total liver volume, and...

Read this Scientific Article Protocol about Author Spotlight: Hepatocellular Carcinoma Research Through Patient-Derived Organoid Engraftment in Liver Regeneration Models at JoVE.com

A Hepatocellular Cancer Patient&#45;Derived Organoid Xenograft Model to Investigate Impact of Liver Regeneration on Tumor Growth

Organoids generated from patient tumors are orthotopically injected into the mouse liver. The resection of non-tumorous liver tissue leads to a regenerative environment in the liver tissue where the tumor is located.

Recurrence poses a notable challenge after hepatocellular carcinoma (HCC) treatment, impacting more than 70% of patients who undergo surgical resection. ...

a-hepatocellular-cancer-patient-derived-organoid-xenograft-model-to

Research

JoVE Journal

Cancer Research

632 ビュー.  University Center for Gastrointestinal and Liver Diseases, St. Clara Hospital and University Hospital Basel. Organoids generated from patient tumors are orthotopically injected into the mouse liver. The resection of non-tumorous liver tissue leads to a regenerative environment in the liver tissue where the tumor is located.

ビデオ: A Hepatocellular Cancer Patient-Derived Organoid Xenograft Model to Investigate Impact of Liver Regeneration on Tumor Growth

Development and Maintenance of a Preclinical Patient Derived Tumor Xenograft Model for the Investigation of Novel Anti-Cancer Therapies

Patient derived tumor xenograft (PDTX) models provide a necessary platform in facilitating anti-cancer drug development prior to human trials. Human tumor pieces are injected subcutaneously into athymic nude mice (immunocompromised, T cell deficient) to create a bank of tumors and subsequently are passaged into different generations of mice in order to maintain these tumors from patients. Importantly, cellular heterogeneity of the original tumor is closely emulated in this model, which provides a more clinically relevant model for evaluation of drug efficacy studies (single agent and combination), biomarker analysis, resistant pathways and cancer stem cell biology. Some limitations of the PDTX model include the replacement of the human stroma with mouse stroma after the first generation in mice, inability to investigate treatment effects on metastasis due to the subcutaneous injections of the tumors, and the lack of evaluation of immunotherapies due to the use of immunocompromised mice. However, even with these limitations, the PDTX model provides a powerful preclinical platform in the drug discovery process.

Utilizing patient-derived tumors in a subcutaneous preclinical model is an excellent way to study the efficacy of novel therapies, predictive biomarker discovery, and drug resistant pathways. This model, in the drug development process, is essential in determining the fate of many novel anti-cancer therapies prior to clinical investigation.

前臨床患者由来の腫瘍新規抗がん療法の調査のための異種移植モデルの開発とメンテナンス

Patient derived tumor xenograft (PDTX) models provide a necessary platform in facilitating anti-cancer drug development prior to human trials. Human tumor ...

がん研究

Study of Viral Vectors in a Three-dimensional Liver Model Repopulated with the Human Hepatocellular Carcinoma Cell Line HepG2

This protocol describes the generation of a three-dimensional (3D) ex vivo liver model and its application to the study and development of viral vector systems. The model is obtained by repopulating the extracellular matrix of a decellularized rat liver with a human hepatocyte cell line. The model permits studies in a vascularized 3D cell system, replacing potentially harmful experiments with living animals. Another advantage is the humanized nature of the model, which is closer to human physiology than animal models.
In this study, we demonstrate the transduction of this liver model with a viral vector derived from adeno-associated viruses (AAV vector). The perfusion circuit that supplies the 3D liver model with media provides an easy means to apply the vector. The system permits monitoring of the major metabolic parameters of the liver. For final analysis, tissue samples can be taken to determine the extent of recellularization by histological techniques. Distribution of the virus vector and expression of the delivered transgene can be analyzed by quantitative PCR (qPCR), Western blotting and immunohistochemistry. Numerous applications of the vector model in basic research and in the development of gene therapeutic applications can be envisioned, including the development of novel antiviral therapeutics, cancer research, and the study of viral vectors and their potential side effects.

The recellularized extracellular matrix of a decellularized rat liver can be used as a humanized, three-dimensional ex vivo model to study the distribution and transgene expression of a virus or viral vector.

ヒト肝細胞癌細胞株HepG2細胞と三次元肝モデルに再増殖におけるウイルスベクターの研究

This protocol describes the generation of a three-dimensional (3D) ex vivo liver model and its application to the study and development of viral vector ...

Tumor Engraftment in a Xenograft Mouse Model of Human Mantle Cell Lymphoma

B lymphocytes are key players in immune cell circulation and they mainly home to and reside in lymphoid organs. While normal B cells only proliferate when stimulated by T lymphocytes, oncogenic B cells survive and expand autonomously in undefined organ niches. Mantle cell lymphoma (MCL) is one such B cell disorder, where the median survival rate of patients is 4 - 5 years. This calls for the need of effective mechanisms by which the homing and engraftment of these cells are blocked in order to increase the survival and longevity of patients. Therefore, the effort to develop a xenograft mouse model to study the efficacy of MCL therapeutics by blocking the homing mechanism in vivo is of utmost importance. Development of animal recipients for human cell xenotransplantation to test early stage drugs have long been pursued, as relevant preclinical mouse models are crucial to screen new therapeutic agents. This animal model is developed to avoid human graft rejection and to establish a model for human diseases, and it may be an extremely useful tool to study disease progression of different lymphoma types and to perform preclinical testing of candidate drugs for hematologic malignancies, like MCL. We established a xenograft mouse model that will serve as an excellent resource to study and develop novel therapeutic approaches for MCL.

Mantle cell lymphoma (MCL) is a difficult to treat B cell disorder and it is equally difficult to establish a xenograft mouse model of primary MCL to study and develop therapeutics. Here, we describe the successful establishment of MCL xenografts in mice to help understand its underlying biology.

人間マントル細胞リンパ腫の異種移植マウス モデルにおける腫瘍生着

B lymphocytes are key players in immune cell circulation and they mainly home to and reside in lymphoid organs. While normal B cells only proliferate when ...

Patient-derived Orthotopic Xenograft Models for Human Urothelial Cell Carcinoma and Colorectal Cancer Tumor Growth and Spontaneous Metastasis

Cancer patients have poor prognoses when lymph node (LN) involvement is present in both high-grade urothelial cell carcinoma (HG-UCC) of the bladder and colorectal cancer (CRC). More than 50% of patients with muscle-invasive UCC, despite curative therapy for clinically-localized disease, will develop metastases and die within 5 years, and metastatic CRC is a leading cause of cancer-related deaths in the US. Xenograft models that consistently mimic UCC and CRC metastasis seen in patients are needed. This study aims to generate patient-derived orthotopic xenograft (PDOX) models of UCC and CRC for primary tumor growth and spontaneous metastases under the influence of LN stromal cells mimicking the progression of human metastatic diseases for drug screening. Fresh UCC and CRC tumors were obtained from consented patients undergoing resection for HG-UCC and colorectal adenocarcinoma, respectively. Co-inoculated with LN stromal cell (LNSC) analog HK cells, luciferase-tagged UCC cells were intra-vesically (IB) instilled into female non-obese diabetic/severe combined immunodeficiency (NOD/SCID) mice, and CRC cells were intra-rectally (IR) injected into male NOD/SCID mice. Tumor growth and metastasis were monitored weekly using bioluminescence imaging (BLI). Upon sacrifice, primary tumors and mouse organs were harvested, weighed, and formalin-fixed for Hematoxylin and Eosin and immunohistochemistry staining. In our unique PDOX models, xenograft tumors resemble patient pre-implantation tumors. In the presence of HK cells, both models have high tumor implantation rates measured by BLI and tumor weights, 83.3% for UCC and 96.9% for CRC, and high distant organ metastasis rates (33.3% detected liver or lung metastasis for UCC and 53.1% for CRC). In addition, both models have zero mortality from the procedure. We have established unique, reproducible PDOX models for human HG-UCC and CRC, which allow for tumor formation, growth, and metastasis studies. With these models, testing of novel therapeutic drugs can be performed efficiently and in a clinically-mimetic manner.

This protocol describes the generation of patient-derived orthotopic xenograft models by intra-vesically instilling high-grade urothelial cell carcinoma cells or intra-rectally injecting colorectal cancer cells into non-obese diabetic/severe combined immunodeficiency (NOD/SCID) mice for primary tumor growth and spontaneous metastases under the influence of lymph node stromal cells, which mimics the progression of human metastatic diseases.

ヒト尿路上皮細胞癌および大腸癌腫瘍増殖および自発性転移に対する患者由来の眼科医異所性Xenograft移植片モデル

Cancer patients have poor prognoses when lymph node (LN) involvement is present in both high-grade urothelial cell carcinoma (HG-UCC) of the bladder and ...

In Vivo Inhibition of MicroRNA to Decrease Tumor Growth in Mice

MicroRNAs (miRNAs) are important regulators of gene expression through their ability to destabilize mRNA and inhibit translation of target mRNAs. An ever-increasing number of studies have identified miRNAs as potential biomarkers for cancer diagnosis and prognosis, and also as therapeutic targets, adding an extra dimension to cancer evaluation and treatment. In the context of thyroid cancer, tumorigenesis results not only from mutations in important genes, but also from the overexpression of many miRNAs. Accordingly, the role of miRNAs in the control of thyroid gene expression is evolving as an important mechanism in cancer. Herein, we present a protocol to examine the effects of miRNA-inhibitor delivery as a therapeutic modality in thyroid cancer using human tumor xenograft and orthotopic mouse models. After engineering stable thyroid tumoral cells expressing GFP and luciferase, cells are injected into nude mice to develop tumors, which can be followed by bioluminescence. The in vivo inhibition of a miRNA can reduce tumor growth and upregulate miRNA gene targets. This method can be used to assess the importance of a determined miRNA in vivo, in addition to identifying new therapeutic targets.

This protocol describes xenograft and orthotopic mouse models of human thyroid tumorigenesis as a platform to test microRNA-based inhibitor treatments. This approach is ideal to study the function of non-coding RNAs and their potential as new therapeutic targets.

マウスの腫瘍増殖を減少させるマイクロRNAの生体内阻害

MicroRNAs (miRNAs) are important regulators of gene expression through their ability to destabilize mRNA and inhibit translation of target mRNAs. An ...

A Melanoma Patient-Derived Xenograft Model

Accumulating evidence suggests that molecular and biological properties differ in melanoma cells grown in traditional two-dimensional tissue culture vessels versus in vivo in human patients. This is due to the bottleneck selection of clonal populations of melanoma cells that can robustly grow in vitro in the absence of physiological conditions. Further, responses to therapy in two-dimensional tissue cultures overall do not faithfully reflect responses to therapy in melanoma patients, with the majority of clinical trials failing to show the efficacy of therapeutic combinations shown to be effective in vitro. Although xenografting of melanoma cells into mice provides the physiological in vivo context absent from two-dimensional tissue culture assays, the melanoma cells used for engraftment have already undergone bottleneck selection for cells that could grow under two-dimensional conditions when the cell line was established. The irreversible alterations that occur as a consequence of the bottleneck include changes in growth and invasion properties, as well as the loss of specific subpopulations. Therefore, models that better recapitulate the human condition in vivo may better predict therapeutic strategies that effectively increase the overall survival of patients with metastatic melanoma. The patient-derived xenograft (PDX) technique involves the direct implantation of tumor cells from the human patient to a mouse recipient. In this manner, tumor cells are consistently grown under physiological stresses in vivo and never undergo the two-dimensional bottleneck, which preserves the molecular and biological properties present when the tumor was in the human patient. Notable, PDX models derived from organ sites of metastases (i.e., brain) display similar metastatic capacity, while PDX models derived from therapy naive patients and patients with acquired resistance to therapy (i.e., BRAF/MEK inhibitor therapy) display similar sensitivity to therapy.

Patient-derived xenograft (PDX) models more robustly recapitulate melanoma molecular and biological features and are more predictive of therapy response compared to traditional plastic tissue culture-based assays. Here we describe our standard operating protocol for the establishment of new PDX models and the characterization/experimentation of existing PDX models.

黒色腫患者由来異種移植片モデル

Accumulating evidence suggests that molecular and biological properties differ in melanoma cells grown in traditional two-dimensional tissue culture ...

Generation of a Liver Orthotopic Human Uveal Melanoma Xenograft Platform in Immunodeficient Mice

In recent decades, subcutaneously implanted patient-derived xenograft tumors or cultured human cell lines have been increasingly recognized as more representative models to study human cancers in immunodeficient mice than traditional established human cell lines in vitro. Recently, orthotopically implanted patient-derived tumor xenograft (PDX) models in mice have been developed to better replicate features of patient tumors. A liver orthotopic xenograft mouse model is expected to be a useful cancer research platform, providing insights into tumor biology and drug therapy. However, liver orthotopic tumor implantation is generally complicated. Here we describe our protocols for the orthotopic implantation of patient-derived liver-metastatic uveal melanoma tumors. We cultured human liver metastatic uveal melanoma cell lines into immunodeficient mice. The protocols can result in consistently high technical success rates using either a surgical orthotopic implantation technique with chunks of patient-derived uveal melanoma tumor or a needle injection technique with cultured human cell line. We also describe protocols for CT scanning to detect interior liver tumors and for re-implantation techniques using cryopreserved tumors to achieve re-engraftment. Together, these protocols provide a better platform for liver orthotopic tumor mouse models of liver metastatic uveal melanoma in translational research.

Orthotopic human liver metastatic uveal melanoma xenograft mouse models were created using surgical orthotopic implantation techniques with patient-derived tumor chunk and needle injection techniques with cultured human uveal melanoma cell lines.

免疫不全マウスにおける肝直交細胞性ヒトうveal黒色腫異種移植プラットフォームの生成

In recent decades, subcutaneously implanted patient-derived xenograft tumors or cultured human cell lines have been increasingly recognized as more ...

A Mouse Model to Investigate the Role of Cancer-Associated Fibroblasts in Tumor Growth

Cancer-associated fibroblasts (CAFs) can play an important role in tumor growth by creating a tumor-promoting microenvironment. Models to study the role of CAFs in the tumor microenvironment can be helpful for understanding the functional importance of fibroblasts, fibroblasts from different tissues, and specific genetic factors in fibroblasts. Mouse models are essential for understanding the contributors to tumor growth and progression in an in vivo context. Here, a protocol in which cancer cells are mixed with fibroblasts and introduced into mice to develop tumors is provided. Tumor sizes over time and final tumor weights are determined and compared among groups. The protocol described can provide more insight into the functional role of CAFs in tumor growth and progression.

A protocol to co-inject cancer cells and fibroblasts and monitor tumor growth over time is provided. This protocol can be used to understand the molecular basis for the role of fibroblasts as regulators of tumor growth.

腫瘍増殖における癌関連線維芽細胞の役割を調べるためのマウスモデル

Cancer-associated fibroblasts (CAFs) can play an important role in tumor growth by creating a tumor-promoting microenvironment. Models to study the role ...

A Murine Ommaya Xenograft Model to Study Direct-Targeted Therapy of Leptomeningeal Disease

Leptomeningeal disease (LMD) is an uncommon type of central nervous system (CNS) metastasis to the cerebral spinal fluid (CSF). The most common cancers that cause LMD are breast and lung cancers and melanoma. Patients diagnosed with LMD have a very poor prognosis and generally survive for only a few weeks or months. One possible reason for the lack of efficacy of systemic therapy against LMD is the failure to achieve therapeutically effective concentrations of drug in the CSF because of an intact and relatively impermeable blood-brain barrier (BBB) or blood-CSF barrier across the choroid plexus. Therefore, directly administering drugs intrathecally or intraventricularly may overcome these barriers. This group has developed a model that allows for the effective delivery of therapeutics (i.e., drugs, antibodies, and cellular therapies) chronically and the repeated sampling of CSF to determine drug concentrations and target modulation in the CSF (when the tumor microenvironment is targeted in mice). The model is the murine equivalent of a magnetic resonance imaging-compatible Ommaya reservoir, which is used clinically. This model, which is affixed to the skull, has been designated as the &#34;Murine Ommaya.&#34; As a therapeutic proof of concept, human epidermal growth factor receptor 2 antibodies (clone 7.16.4) were delivered into the CSF via the Murine Ommaya to treat mice with LMD from human epidermal growth factor receptor 2-positive breast cancer. The Murine Ommaya increases the efficiency of drug delivery using a miniature access port and prevents the wastage of excess drug; it does not interfere with CSF sampling for molecular and immunological studies. The Murine Ommaya is useful for testing novel therapeutics in experimental models of LMD.

Here, we describe a murine xenograft model that functionally resembles an Ommaya reservoir in patients. We developed the Murine Ommaya to study novel therapeutics for the universally fatal leptomeningeal disease.

レプトメニンゲレス病の直接標的療法を研究するマウス・オンマヤ異種移植片モデル

Leptomeningeal disease (LMD) is an uncommon type of central nervous system (CNS) metastasis to the cerebral spinal fluid (CSF). The most common cancers ...

An Ex Vivo Brain Slice Model to Study and Target Breast Cancer Brain Metastatic Tumor Growth

Brain metastasis is a serious consequence of breast cancer for women as these tumors are difficult to treat and are associated with poor clinical outcomes. Preclinical mouse models of breast cancer brain metastatic (BCBM) growth are useful but are expensive, and it is difficult to track live cells and tumor cell invasion within the brain parenchyma. Presented here is a protocol for ex vivo brain slice cultures from xenografted mice containing intracranially injected breast cancer brain-seeking clonal sublines. MDA-MB-231BR luciferase tagged cells were injected intracranially into the brains of Nu/Nu female mice, and following tumor formation, the brains were isolated, sliced, and cultured ex vivo. The tumor slices were imaged to identify tumor cells expressing luciferase and monitor their proliferation and invasion in the brain parenchyma for up to 10 days. Further, the protocol describes the use of time-lapse microscopy to image the growth and invasive behavior of the tumor cells following treatment with ionizing radiation or chemotherapy. The response of tumor cells to treatments can be visualized by live-imaging microscopy, measuring bioluminescence intensity, and performing histology on the brain slice containing BCBM cells. Thus, this ex vivo slice model may be a useful platform for rapid testing of novel therapeutic agents alone or in combination with radiation to identify drugs personalized to target an individual patient&#39;s breast cancer brain metastatic growth within the brain microenvironment.

An Ex Vivo Brain Slice Model to Study and Target Breast Cancer Brain Metastatic Tumor Growth

We introduce a protocol for measuring real-time drug and radiation response of breast cancer brain metastatic cells in an organotypic brain slice model. The methods provide a quantitative assay to investigate the therapeutic effects of various treatments on brain metastases from breast cancer in an ex vivo manner within the brain microenvironment interface.