This work was supported by a Canadian Institutes of Health Research Phase 1 Clinician-Scientist Award (A.G.) and an Operating Grant from the Cancer Research Society (A.G.). The authors are grateful to Dr. John Dick for his support of this project.

A schematic overview of the protocol is presented in Figure 1.
1. Processing of Human HCC Samples
Obtain primary human HCC specimens with written patient consent and with the approval of the institutional research ethics board. These protocols have been carried out at our institution with approval from the University Health Network Research Ethics Board in compliance with all institutional, national, and international guidelines for human welfare.
Collect fresh HCC specimens as soon as possible following the surgical procedure once appropriate samples have been taken for clinical purposes. Ideally this should take place within 30 min after removal of the tissue from the patient. As illustrated in Figure 2, a sample of at least 1 cm3 obtained from the periphery of the tumor is optimal, as the central portion of the tumor may be necrotic. Tumors that have not received any treatment prior to resection such as radiation, chemotherapy, or ablation are preferred in order to maximize the chance that tumor cells are viable. Handle primary human tissues in accordance with standard personal protective protocols for biohazardous material. Perform all laboratory manipulations of tumor tissues and cell preparations in a class II biosafety cabinet using aseptic techniques.
<ol>
	<li>Place the fresh HCC sample in 10-25 ml serum-free Dulbecco&#39;s Modified Eagle Medium/Ham&#39;s F12 at 4 &#176;C and transfer on ice to the laboratory for immediate processing and preparation of tissue fragments and/or cells for xenografting into mice.</li>
	<li>Using sterile forceps, place the tumor sample in a 100 mm x 20 mm Petri dish or other suitable sterile working surface. Divide the tumor sample into fragments of approximately 2-3 mm3 using a No.10 surgical scalpel blade. At this point, consider snap-freezing or formalin-fixing some tumor fragments for other experiments or analyses as required.</li>
	<li>For xenografting of tumor fragments, place some of the HCC fragments into one or more microcentrifuge tubes containing enough Matrigel to allow the fragments to remain submerged. Keep these tubes on ice.</li>
	<li>For preparation of a tumor cell suspension, use the surgical scalpel blade to mince the remaining HCC tissue as much as possible and mix with 5-10 ml of DMEM-F12 in a 50 ml conical tube depending on the volume of the minced tissue.</li>
	<li>Add collagenase type IV and dispase II at final concentrations of 200 units/ml and 0.8 units/ml respectively. Pipette the mixture up and down well using a 25 ml pipette.</li>
	<li>Seal the tube and incubate the mixture from step 1.6 at 37 &#176;C in a 5% carbon dioxide incubator for 30-60 min depending on the softness of the tumor tissue. Pipette the mixture up and down a few times every 10 min to assess the progress of the enzymatic digestion.</li>
	<li>After digestion is complete, pass the tumor solution through a 100 &#956;m cell strainer. Gently mash the remaining tissue on the cell strainer using a 25 ml pipette tip to enable the maximum number of tumor cells to pass through. Collect the strained cell suspension in a 15 ml conical tube.</li>
	<li>Centrifuge the tumor cell suspension at 1,200 rpm for 5 min at 4 &#176;C.</li>
	<li>Gently decant the supernatant. Depending on the size of the pellet, add 2-5 ml of ice cold 1x red blood cell (RBC) lysis buffer and gently pipette up and down to resuspend the pellet. Keep on ice for 5 min.</li>
	<li>Add DMEM-F12 to a total volume of 15 ml and centrifuge at 1,000 rpm for 5 min at 4 &#176;C to wash out the RBC lysis buffer.</li>
	<li>Gently decant the supernatant and resuspend the RBC-free tumor cell pellet in DMEM-F12.</li>
	<li>Count the viable cells using trypan blue exclusion either manually or with an automated cell counter.</li>
	<li>Centrifuge tumor cell aliquots containing the desired number of cells for injection as described above, resuspend the resulting cell pellet in 30 &#956;l of ice-cold Matrigel, and store on ice.</li>
</ol>
Optional: After step 1.11, we routinely deplete human CD45+ cells (leukocytes) from the bulk tumor cell suspension and/or purify subsets of tumor cells using flow cytometry or immunomagnetic beads. Detailed protocols for these techniques are well described by the manufacturers of the relevant antibodies, beads, and flow cytometers.
Note: The protocol described above can also be used to process human tumor xenografts harvested from mice in order to perform serial transplantation, substituting the xenograft tissue for the primary human HCC tissue in step 1.1. In this situation, after step 1.11, we routinely deplete infiltrating murine cells from the cell suspension using an antibody against the mouse histocompatibility antigen H2k.
2. Xenografting
Conduct all animal procedures in compliance with protocols approved by the institutional animal care committee. The procedures described herein have been completed under a specific Animal Use Protocol approved by the University Health Network Animal Care Committee in accordance and compliance with all relevant regulatory and institutional agencies, regulations, and guidelines.
Equipment for delivery of inhalational volatile anesthetic agents to small animals should be utilized according to standard operating procedures of the animal facility and research institute. Carry out all surgical procedures using aseptic technique and sterile instruments in a class II biosafety cabinet. Utilize non-obese diabetic severe combined immunodeficiency (NOD/SCID) or NOD/SCID/interleukin 2 receptor gamma chain null (NSG) strains of mice of either sex at 6-8 weeks of age (The Jackson Laboratory, Bar Harbor, ME) 9,10. These mice must be housed and maintained in a facility capable of providing pathogen-free conditions suitable for immunodeficient animals.
Prepare mice for surgery in a chamber supplying 5% (v/v) inhaled Isoflurane in 1 L/min of oxygen. Maintain anesthesia until there is loss of corneal and toe reflex in the animal(s). For subcutaneous xenografting, shave one or more small areas on the dorsum of the animal(s) and cleanse the skin with 70% ethanol. For intrahepatic xenografting, shave the ventral thorax and abdomen of the animal(s) from the axillae down to the inguinal region and cleanse the skin with 70% ethanol.
2.1 Subcutaneous Implantation of Tumor Fragments
<ol>
	<li>Place the anesthetized mouse prone, maintaining inhalational Isoflurane anesthesia (2% (v/v) in 1 L/min O2) with a mouthpiece. Apply Tear-Gel to protect the animal&#39;s eyes from traumatic injury.</li>
	<li>Sterilize the dorsal shaved area(s) with Betadine surgical scrub followed by 70% ethanol and lastly with povidone-iodine solution.</li>
	<li>Make a 5 mm skin incision using sterile sharp scissors.</li>
	<li>Insert a closed blunt scissor gently into the subcutaneous space and spread gently to develop a pocket large enough to accommodate a tumor fragment.</li>
	<li>Insert the tumor fragment prepared in step 1.3 into the subcutaneous pocket using sterile fine forceps.</li>
	<li>Close the skin incision using sutures or clips.</li>
	<li>Provide postoperative care to the mouse as described below.</li>
</ol>
2.2 Subcutaneous injection of tumor cells
<ol>
	<li>Prepare the animal as described in steps 2.1.1 and 2.1.2.</li>
	<li>Load the suspension of tumor cells in Matrigel prepared in step 1.13 into an insulin syringe with a 29 G 1/2 in needle.</li>
	<li>Insert the needle into the subcutaneous space and discharge the contents of the syringe. Advancing the needle several millimetres along the subcutaneous plane away from the skin puncture site prevents leakage of the tumor cell suspension out of the puncture site upon withdrawal of the needle.</li>
	<li>Provide postoperative care to the mouse as described below.</li>
</ol>
2.3 Intrahepatic Implantation of Tumor Fragments
<ol>
	<li>Using a 27 G 1/2 in needle on a 1 ml syringe, administer 350 &#956;l of sterile normal saline solution subcutaneously in the dorsum of the anesthetized animal's neck to compensate for intraoperative fluid losses. For analgesia, administer 350 &#956;l of sterile normal saline containing buprenorphine (0.1 mg/kg) subcutaneously on the animal&#39;s flank, using a 27 G 1/2-in needle on a 1 ml syringe.</li>
	<li>Place the mouse supine on a pre-heated pad with the nose and mouth positioned inside the mouthpiece to deliver maintenance inhalational Isoflurane anesthesia (2% (v/v) in 1 L/min O2).</li>
	<li>Extend the limbs and secure them with tape to the operating surface to optimize exposure of the ventral abdomen and thorax.</li>
	<li>Perform the procedure under a magnifying lamp to optimize visualization.</li>
	<li>Sterilize the shaved skin on the ventral abdomen and thorax with Betadine surgical scrub followed by 70% ethanol and lastly with povidone-iodine solution.</li>
	<li>Using sterile sharp scissors, make a transverse bilateral subcostal skin incision and divide the muscle layers to enter the peritoneal cavity to allow adequate exposure of the entire liver.</li>
	<li>Place a stitch in the skin above the xiphoid process and secure it to the mouthpiece with tape in order to allow better exposure of the liver and surrounding structures.</li>
	<li>Use two cotton-tipped applicators adjacent and posterior to the liver to stabilize it.</li>
	<li>Make an incision 3 mm in length and depth on the surface of the liver using a sterile No. 10 scalpel blade.</li>
	<li>Immediately apply Surgicel and gentle pressure to the incision site to achieve hemostasis; remove after 60-90 sec and proceed only if complete hemostasis has been achieved.</li>
	<li>Place a tumor fragment step 1.3 into the liver incision with sterile fine forceps or with an 18 G needle.</li>
	<li>Apply a small piece of Surgicel over the liver incision to prevent displacement of the tumor fragment and ensure continued hemostasis.</li>
	<li>Close the incision with sutures or clips.</li>
	<li>Provide postoperative care as described below</li>
</ol>
2.4 Intrahepatic Implantation of Tumor Cells via Direct Injection into the Liver
<ol>
	<li>Prepare the mouse as described in Steps 2.3.1 to 2.3.7 above.</li>
	<li>Load the tumor cell suspension (prepared in step 1.13) into an insulin syringe using a 29 G 1/2 in needle.</li>
	<li>With the liver stabilized using a cotton-tip applicator, insert the insulin syringe needle into the liver and advance the tip a few millimetres beyond the puncture site along a subcapsular plane.</li>
	<li>Gently discharge the contents of the syringe and then remove the needle from the liver.</li>
	<li>Place Surgicel over the puncture site and apply gentle pressure with a cotton-tipped applicator to prevent leakage of the tumor cell suspension and to achieve complete hemostasis.</li>
	<li>Close the incision with sutures or clips and provide postoperative care as described below.</li>
</ol>
2.5 Intrahepatic Xenografting of Tumor Cells via Injection into the Spleen
<ol>
	<li>Prepare the mouse as described in steps 2.3.1 to 2.3.5 above.</li>
	<li>Using sterile sharp scissors, make a 1 cm left subcostal incision to enter the peritoneal cavity.</li>
	<li>Use a cotton-tipped applicator to reflect the stomach cranially and to the animal&#39;s right side in order to expose the spleen.</li>
	<li>By handling the surrounding adipose tissues with sterile fine atraumatic forceps, deliver the spleen into the incision and place a cotton-tipped applicator behind the spleen to stabilize it.</li>
	<li>Using 5-0 silk suture, place a loose pre-tied knot around the spleen above the lower pole.</li>
	<li>Load the tumor cell suspension (prepared in step 1.13) into an insulin syringe using a 29 G 1/2 in needle.</li>
	<li>Insert the insulin syringe needle into the lower pole of the spleen and advance it past the level of the loose pre-tied knot.</li>
	<li>Slowly discharge the contents of the syringe, remove the needle from the spleen, and tighten the knot to prevent any leakage of the injected tumor cell suspension.</li>
	<li>Replace the spleen into the peritoneal cavity.</li>
	<li>Close the incision with sutures or clips and provide postoperative care as described below.</li>
</ol>
2.6 Partial Hepatectomy to Facilitate Intrahepatic Engraftment of Human Tumor Tissue
<ol>
	<li>Prepare the mouse as described in steps 2.3.1 to 2.3.7.</li>
	<li>With sterile sharp scissors, divide the falciform ligament attached to the median lobe of the liver.</li>
	<li>Using a cotton-tipped applicator, mobilize the left lobe of the liver and position a loose pre-tied knot of 5-0 silk suture around the left lobe. Advance the loose knot as close as possible towards the bilio-vascular pedicle of the left lobe and tighten the knot.</li>
	<li>Excise the left lobe distal to the ligature using sterile scissors, leaving a small stump to prevent slippage of the knot and subsequent hemorrhage.</li>
	<li>Using a similar technique, ligate and resect the majority of the median lobe of the liver, avoiding the gallbladder.</li>
	<li>Use Surgicel and gentle pressure with a cotton-tipped applicator along the cut surfaces of the liver parenchyma to achieve complete hemostasis.</li>
	<li>For intrahepatic xenografting of a tumor fragment or tumor cells, proceed with steps 2.3.8 to 2.3.11 or 2.4.2 to 2.4.5 as described above.</li>
	<li>For intrahepatic xenografting of tumor cells via splenic injection, utilize cotton-tipped applicators to gently reflect the stomach cranially and to the animal's right side, exposing the spleen. Proceed with steps 2.5.4 to 2.5.9 as described above.</li>
	<li>Close the incision with sutures or clips and provide postoperative care as described below.</li>
</ol>
2.7 Postoperative Care
<ol>
	<li>Remove the mouse from the inhalational anesthesia mouthpiece.</li>
	<li>Place the mouse in a cage under a heat lamp for approximately 20 min until recovered from anesthesia and mobilizing fully.</li>
	<li>Repeat the buprenorphine dose every 8-12 hr during the first 2-3 postoperative days.</li>
</ol>

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The authors declare that they have no competing financial interests.

We have described a variety of techniques to establish subcutaneous and intrahepatic human HCC xenografts in immunodeficient mice that can be applied to a wide variety of experimental questions and assays. While subcutaneous xenografts have been widely used to study various aspects of HCC biology, intrahepatic xenografts are rarely described in the literature. Furthermore, the majority of studies describing the use of xenografts have generated these from well established cell lines. Given the limitations of cancer cell l...

Hepatocellular carcinoma (HCC) is the fifth most common cancer worldwide and the most rapidly increasing cause of cancer death in North America. The most prevalent risk factor for HCC is liver cirrhosis, most frequently occurring due to chronic viral hepatitis, alcohol misuse, autoimmune disease, or hereditary metabolic disorders 1.
Despite the heavy disease burden imposed by HCC on populations worldwide, the pathophysiology of HCC is relatively poorly understood in comparison to other common cancers such as colorectal, breast, or prostate cancer. For example, specific molecular and cellular events driving tumorigenesis remain to be clearly defined 2. Like most other solid epithelial cancers, genomic approaches have revealed heterogeneity in the aberrations associated with HCC 3. A number of studies have revealed disordered activity of a variety of signaling pathways involved in cell proliferation, survival, differentiation, and angiogenesis 4. In addition, the role of cancer stem cells in HCC pathobiology remains to be clarified 5.
With a limited understanding of HCC pathophysiology, the armamentarium of effective therapies for HCC has also remained relatively limited. Early-stage patients with tumors confined to the liver are candidates for curative therapy using tumor ablation or surgical resection, though recurrence is common. For patients with more advanced disease, chemotherapy and radiation are of limited efficacy and are used primarily for disease control with palliative intent 6.
High quality in vivo experimental models of human HCC thus provide a valuable platform for much needed basic research into the pathophysiology of human HCC as well as for evaluation of novel therapeutic approaches. As compared with the use of cell lines or highly defined mouse models, xenografts of primary human tumors in immunodeficient mice have emerged as valuable tools for such studies since they are capable of recapitulating the human disease with high fidelity while also capturing the heterogeneity that is present within and between different patients 7,8. To this end, we have developed a variety of methods to establish human HCC xenografts in immunodeficient mice. While the majority of published studies involving HCC xenografts describe the use of well-established human HCC cell lines for this purpose, we have focused on optimizing our assays to generate xenografts from primary HCC specimens obtained immediately after surgical resection from patients.
Different xenografting techniques may be required for different research applications. For example, subcutaneous xenografts generated from tumor fragments are generated rapidly, are easily monitored, and may be more appropriate for local administration of novel therapeutics with convenient monitoring of tumor response. Intrahepatic xenografts may be more relevant for studies pertaining to the role of the hepatic microenvironment in HCC biology. Xenografts generated from tumor cell suspensions are necessary for the identification and characterization of tumor-initiating cell subsets or for experiments requiring in vitro manipulations of tumor cells prior to xenotransplantation. We have thus developed and validated the following protocols to establish subcutaneous or intrahepatic xenografts from cell suspensions or tumor fragments derived from primary human HCC specimens.

<table border="1">
	<tbody>
		<tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Dulbecco's Mod. Eagle Medium/Ham's F12 50/50 Mix x1(DMEM-F12)</td>
			<td>WISENT Bioproducts</td>
			<td>319-075-CL</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Collagenase TypeIV</td>
			<td>Sigma-Aldrich</td>
			<td>C5138</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dispase II</td>
			<td>Stemcell Technologies</td>
			<td>7923</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Matrigel Matrix</td>
			<td>Becton-Dickinson Biosciences</td>
			<td>354234</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>10 % Buffered Formalin solution</td>
			<td>Sigma-Aldrich</td>
			<td>HT501128</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>0.9 % Saline Solution (NaCl), sterile</td>
			<td>House Brand</td>
			<td>1011-L8001</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Betadine surgical scrub</td>
			<td>Purdue Pharma</td>
			<td>NPN 00158313</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>LORIS 10% PVP-I Solution</td>
			<td>LERNA Pharma Inc.</td>
			<td>109-09</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Buprenorphine (Temegesic) NR 0.3 mg/ml</td>
			<td>Reckitt Benckiser</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Isoflurane USP, 99.9 %, inhalation anesthetic</td>
			<td>Pharmaceutical Partners of Canada Inc.</td>
			<td>M60302</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tear-Gel</td>
			<td>Novartis Pharmaceuticals</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Frozen section compound</td>
			<td>VWR</td>
			<td>95057-838</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cryomold, Tissue -Tek</td>
			<td>Sakura Finetek</td>
			<td>4566</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Precision Glide Needle 18G 1 &frac12;</td>
			<td>Becton-Dickinson Biosciences</td>
			<td>305196</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Precision Glide Needle 27G &frac12;</td>
			<td>Becton-Dickinson Biosciences</td>
			<td>305109</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Insulin syringe, 3/10 cc U-100, 29G&frac12;</td>
			<td>Becton-Dickinson Biosciences</td>
			<td>309301</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Surgical blade No.10</td>
			<td>Feather Safety Razor Co.</td>
			<td>08-916-5A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>#5-0 Soft silk surgical suture, 3/8" taper point needle</td>
			<td>Syneture</td>
			<td>VS-880</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Transpore surgical tape</td>
			<td>3M Health care</td>
			<td>1577-1</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cotton applicator</td>
			<td>Medpro</td>
			<td>018-425</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Surgicel, oxidized regenerated cellulose</td>
			<td>Ethicon</td>
			<td>1951</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Cell strainer 100 &mu;m nylon</td>
			<td>Becton-Dickinson Biosciences</td>
			<td>352360</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Magnification lighting with mobile base</td>
			<td>Benson medical Industries Inc.</td>
			<td>model: RLM-CLT-120V</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Petridish sterile 100x20 mm</td>
			<td>Sarstedt</td>
			<td>821474</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tissue forcep, 1x2 teeth, 4-1/2"</td>
			<td>Almedic</td>
			<td>A10-302</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Adson dressing forcep 4-3/4"</td>
			<td>Almedic</td>
			<td>A10-220</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Eye dressing forcep, serrated, straight, 4"</td>
			<td>Almedic</td>
			<td>A19-560</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hartman Hemostatic Forceps, curved, 3-1/2"</td>
			<td>Almedic</td>
			<td>A12-142</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Iris scissor, curved, 4-1/4"</td>
			<td>Almedic</td>
			<td>A8-690</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Iris scissor, straight, 4-1/2"</td>
			<td>Almedic</td>
			<td>A8-684</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Olsen-Hegan needle driver, 5-1/2"</td>
			<td>Almedic</td>
			<td>A17-228</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

generation of subcutaneous and intrahepatic human hepatocellular carcinoma xenografts in immunodeficient mice

In vivo experimental models of hepatocellular carcinoma (HCC) that recapitulate the human disease provide a valuable platform for research into disease pathophysiology and for the preclinical evaluation of novel therapies. We present a variety of methods to generate subcutaneous or orthotopic human HCC xenografts in immunodeficient mice that could be utilized in a variety of research applications. With a focus on the use of primary tumor tissue from patients undergoing surgical resection as a starting point, we describe the preparation of cell suspensions or tumor fragments for xenografting. We describe specific techniques to xenograft these tissues i) subcutaneously; or ii) intrahepatically, either by direct implantation of tumor cells or fragments into the liver, or indirectly by injection of cells into the mouse spleen. We also describe the use of partial resection of the native mouse liver at the time of xenografting as a strategy to induce a state of active liver regeneration in the recipient mouse that may facilitate the intrahepatic engraftment of primary human tumor cells. The expected results of these techniques are illustrated. The protocols described have been validated using primary human HCC samples and xenografts, which typically perform less robustly than the well-established human HCC cell lines that are widely used and frequently cited in the literature. In comparison with cell lines, we discuss factors which may contribute to the relatively low chance of primary HCC engraftment in xenotransplantation models and comment on technical issues that may influence the kinetics of xenograft growth. We also suggest methods that should be applied to ensure that xenografts obtained accurately resemble parent HCC tissues.

Figure 3 demonstrates the typical appearance of a subcutaneous human HCC xenograft and the corresponding histopathological appearance of the tumor. The development and growth of subcutaneous xenografts can be readily monitored by daily examination of recipient mice. The time interval between xenografting and development of a tumor may vary greatly depending on the type of tissue (tumor fragment vs. cell suspension), source of tissue (primary patient sample, passaged xenograft, or cell line), and quantity...

Watch this Scientific Journal Video about Generation of Subcutaneous and Intrahepatic Human Hepatocellular Carcinoma Xenografts in Immunodeficient Mice at JoVE.com

Generation of Subcutaneous and Intrahepatic Human Hepatocellular Carcinoma Xenografts in Immunodeficient Mice

Human tumor xenografts in immunodeficient mice are valuable tools to study cancer biology. Specific protocols to generate subcutaneous and intrahepatic xenografts from human hepatocellular carcinoma cells or tumor fragments are described. Liver regeneration induced by partial hepatectomy in recipient mice is presented as a strategy to facilitate intrahepatic engraftment.

In  vivo experimental models of hepatocellular carcinoma (HCC) that recapitulate  the human disease provide a valuable platform for research into disease  ...

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35.7K Views.  University Health Network. Human tumor xenografts in immunodeficient mice are valuable tools to study cancer biology. Specific protocols to generate subcutaneous and intrahepatic xenografts from human hepatocellular carcinoma cells or tumor fragments are described. Liver regeneration induced by partial hepatectomy in recipient mice is presented as a strategy to facilitate intrahepatic engraftment.

Video: Generation of Subcutaneous and Intrahepatic Human Hepatocellular Carcinoma Xenografts in Immunodeficient Mice

Mammary Transplantation of Stromal Cells and Carcinoma Cells in C57BL/6J Mice

The influence of stromal cells, including fibroblasts on mammary tumor progression has been well documented through the use of mouse models, in particular through transplantation of stromal cells and epithelial cells in the mammary gland of mice. Current transplantation models often involve the use of immunocompromised mice due to the different genetic backgrounds of stromal cells and epithelial cells. Extracellular matrices are often used to embed the two different cell types for consistent cell-cell interactions, but involve the use of Matrigel or rat tail collagen, which are immunogenic substrates. The lack of functional T cells from immunocompromised mice prevents accurate assessment of stromal cells on mammary tumor progression in vivo, with important implications on drug development and efficacy. Moreover, immunocompromised mice are costly, hard to breed and require special care conditions. To overcome these obstacles, we have developed an approach to orthotopically transplant stromal cell and epithelial cells into mice from the same genetic background to induce consistent tumor formation. This system involves harvesting normal, carcinoma associated fibroblasts, PyVmT mammary carcinoma cells and collagen from donor C57BL/6J mice. The cells are then embedded in collagen and transplanted in the inguinal mammary glands of female C57BL/6J mice. Transplantation of PyVmT cells alone form palpable tumors 30-40 days post transplantation. Endpoint analysis at 60 days indicates that co-transplantation with fibroblasts enhances mammary tumor growth compared to PyVmT cells transplanted alone. While cells and matrix from C57BL/6J mice were used in these studies, the isolation of cells and matrix and transplantation approach may be applied towards mice from different genetic backgrounds demonstrating versatility. In summary, this system may be used to investigate molecular interactions between stromal cells and epithelial cells, and overcomes critical limitations in immunocompromised mouse models.

In this report, we demonstrate a system to isolate and culture donor cells from the mouse mammary gland, and orthotopically transplant these cells in recipient mice to analyze stromal: epithelial interactions during mammary tumor development.

The influence of stromal cells, including fibroblasts on mammary tumor progression has been well documented through the use of mouse models, in particular ...

Experimental Generation of Carcinoma-Associated Fibroblasts (CAFs) from Human Mammary Fibroblasts

Carcinomas are complex tissues comprised of neoplastic cells and a non-cancerous compartment referred to as the 'stroma'. The stroma consists of extracellular matrix (ECM) and a variety of mesenchymal cells, including fibroblasts, myofibroblasts, endothelial cells, pericytes and leukocytes 1-3.
The tumour-associated stroma is responsive to substantial paracrine signals released by neighbouring carcinoma cells. During the disease process, the stroma often becomes populated by carcinoma-associated fibroblasts (CAFs) including large numbers of myofibroblasts. These cells have previously been extracted from many different types of human carcinomas for their in vitro culture. A subpopulation of CAFs is distinguishable through their up-regulation of &alpha;-smooth muscle actin (&alpha;-SMA) expression4,5. These cells are a hallmark of 'activated fibroblasts' that share similar properties with myofibroblasts commonly observed in injured and fibrotic tissues 6. The presence of this myofibroblastic CAF subset is highly related to high-grade malignancies and associated with poor prognoses in patients.
Many laboratories, including our own, have shown that CAFs, when injected with carcinoma cells into immunodeficient mice, are capable of substantially promoting tumourigenesis 7-10. CAFs prepared from carcinoma patients, however, frequently undergo senescence during propagation in culture limiting the extensiveness of their use throughout ongoing experimentation. To overcome this difficulty, we developed a novel technique to experimentally generate immortalised human mammary CAF cell lines (exp-CAFs) from human mammary fibroblasts, using a coimplantation breast tumour xenograft model.
In order to generate exp-CAFs, parental human mammary fibroblasts, obtained from the reduction mammoplasty tissue, were first immortalised with hTERT, the catalytic subunit of the telomerase holoenzyme, and engineered to express GFP and a puromycin resistance gene. These cells were coimplanted with MCF-7 human breast carcinoma cells expressing an activated ras oncogene (MCF-7-ras cells) into a mouse xenograft. After a period of incubation in vivo, the initially injected human mammary fibroblasts were extracted from the tumour xenografts on the basis of their puromycin resistance 11.
We observed that the resident human mammary fibroblasts have differentiated, adopting a myofibroblastic phenotype and acquired tumour-promoting properties during the course of tumour progression. Importantly, these cells, defined as exp-CAFs, closely mimic the tumour-promoting myofibroblastic phenotype of CAFs isolated from breast carcinomas dissected from patients. Our tumour xenograft-derived exp-CAFs therefore provide an effective model to study the biology of CAFs in human breast carcinomas. The described protocol may also be extended for generating and characterising various CAF populations derived from other types of human carcinomas.

Experimental Generation of Carcinoma-Associated Fibroblasts CAFs from Human Mammary Fibroblasts

Carcinoma-associated fibroblasts (CAFs) rich in myofibroblasts present within the tumour stroma, play a major role in driving tumour progression. We developed a coimplantation tumour xengraft model for experimentally generating CAFs from human mammary fibroblasts. The protocol describes how to establish CAF myofibroblasts that acquire an ability to promote tumourigenesis.

Carcinomas are complex tissues comprised of neoplastic cells and a non-cancerous compartment referred to as the 'stroma'. The stroma consists of ...

Induction of Invasive Transitional Cell Bladder Carcinoma in Immune Intact Human MUC1 Transgenic Mice:  A Model for Immunotherapy Development

A preclinical model of invasive bladder cancer was developed in human mucin 1 (MUC1) transgenic (MUC1.Tg) mice for the purpose of evaluating immunotherapy and/or cytotoxic chemotherapy. To induce bladder cancer, C57BL/6 mice (MUC1.Tg and wild type) were treated orally with the carcinogen N-butyl-N-(4-hydroxybutyl)nitrosamine (OH-BBN) at 3.0 mg/day, 5 days/week for 12 weeks. To assess the effects of OH-BBN on serum cytokine profile during tumor development, whole blood was collected via submandibular bleeds prior to treatment and every four weeks. In addition, a MUC1-targeted peptide vaccine and placebo were administered to groups of mice weekly for eight weeks. Multiplex fluorometric microbead immunoanalyses of serum cytokines during tumor development and following vaccination were performed. At termination, interferon gamma (IFN-&#947;)/interleukin-4 (IL-4) ELISpot analysis for MUC1 specific T-cell immune response and histopathological evaluations of tumor type and grade were performed. The results showed that: (1) the incidence of bladder cancer in both MUC1.Tg and wild type mice was 67%; (2) transitional cell carcinomas (TCC) developed at a 2:1 ratio compared to squamous cell carcinomas (SCC); (3) inflammatory cytokines increased with time during tumor development; and (4) administration of the peptide vaccine induces a Th1-polarized serum cytokine profile and a MUC1 specific T-cell response. All tumors in MUC1.Tg mice were positive for MUC1 expression, and half of all tumors in MUC1.Tg and wild type mice were invasive. In conclusion, using a team approach through the coordination of the efforts of pharmacologists, immunologists, pathologists and molecular biologists, we have developed an immune intact transgenic mouse model of bladder cancer that expresses hMUC1.

An N-butyl-N-(4-hydroxybutyl)nitrosamine-induced bladder cancer model was developed in human mucin 1 (MUC1) transgenic mice for the purpose of testing MUC1-directed immunotherapy. After administering a MUC1-targeted peptide vaccine, a cytotoxic T lymphocyte response to MUC1 was confirmed by measuring serum cytokine levels and T-cell specific activity.

A preclinical model of invasive bladder cancer was developed in human mucin 1 (MUC1) transgenic (MUC1.Tg) mice for the purpose of evaluating immunotherapy ...

Creating Anatomically Accurate and Reproducible Intracranial Xenografts of Human Brain Tumors

Orthotopic tumor models are currently the best way to study the characteristics of a tumor type, with and without intervention, in the context of a live animal &#8211;&#160;particularly in sites with unique physiological and architectural qualities such as the brain. In vitro and ectopic models cannot account for features such as vasculature, blood brain barrier, metabolism, drug delivery and toxicity, and a host of other relevant factors. Orthotopic models have their limitations too, but with proper technique tumor cells of interest can be accurately engrafted into tissue that most closely mimics conditions in the human brain. By employing methods that deliver precisely measured volumes to accurately defined locations at a consistent rate and pressure, mouse models of human brain tumors with predictable growth rates can be reproducibly created and are suitable for reliable analysis of various interventions. The protocol described here focuses on the technical details of designing and preparing for an intracranial injection, performing the surgery, and ensuring successful and reproducible tumor growth and provides starting points for a variety of conditions that can be customized for a range of different brain tumor models.

The brain is a unique site with qualities that are not well represented by in vitro or ectopic analyses. Orthotopic mouse models with reproducible location and growth characteristics can be reliably created with intracranial injections using a stereotaxic fixation instrument and a low pressure syringe pump.

Orthotopic tumor models are currently the best way to study the characteristics of a tumor type, with and without intervention, in the context of a live ...

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 ...

Th17 Inflammation Model of Oropharyngeal Candidiasis in Immunodeficient Mice

Oropharyngeal Candidiasis (OPC) disease is caused not only due to the lack of host immune resistance, but also the absence of appropriate regulation of infection-induced immunopathology. Although Th17 cells are implicated in antifungal defense, their role in immunopathology is unclear. This study presents a method for establishing oral Th17 immunopathology associated with oral candidal infection in immunodeficient mice. The method is based on reconstituting lymphopenic mice with in vitro cultured Th17 cells, followed by oral infection with Candida albicans (C. albicans). Results show that unrestrained Th17 cells result in inflammation and pathology, and is associated with several measurable read-outs including weight loss, pro-inflammatory cytokine production, tongue histopathology and mortality, showing that this model may be valuable in studying OPC immunopathology. Adoptive transfer of regulatory cells (Tregs) controls and reduces the inflammatory response, showing that this model can be used to test new strategies to counteract oral inflammation. This model may also be applicable in studying oral Th17 immunopathology in general in the context of other oral diseases.

Although Candida infection models are available to study host immune resistance, a model to study T cell mediated immunopathology in the context of Candida infection is absent. Here we describe a method to establish Th17 immunopathology associated with oral Candida infection in immunodeficient mice.

Oropharyngeal Candidiasis (OPC) disease is caused not only due to the lack of host immune resistance, but also the absence of appropriate regulation of ...

A "Patient-Like" Orthotopic Syngeneic Mouse Model of Hepatocellular Carcinoma Metastasis

The majority of cancer-related deaths are caused by the metastasis of the cancer rather than the primary tumor itself. Yet, the underlying mechanisms of cancer metastasis are still unclear. Animal models are essential for elucidating the mechanisms and for evaluating novel strategies for the treatment of metastatic cancers. Here, an in-depth description of a &#8220;patient-like&#8221; orthotopic syngeneic mouse model for exploring the mechanisms of metastasis of solid organ tumors is provided. The survival surgical implantation of BNL 1ME A.7R.1 mouse hepatocellular carcinoma cells directly into the liver (the organ of origin) of the inbred wild-type immune competent laboratory mouse strain, BALB/c is described. The success and reproducibility of this methodology recommends it for widespread use in elucidating the biological mechanisms of solid organ cancer metastasis.

A &quot;Patient-Like&quot; Orthotopic Syngeneic Mouse Model of Hepatocellular Carcinoma Metastasis

The metastatic spread of cancer is the major cause of cancer-related deaths. We provide an in-depth description of our survival surgery methodology for establishing a &#8220;patient-like&#8221; orthotopic syngeneic mouse model system for studying the mechanisms of metastasis in solid organ tumors.

The majority of cancer-related deaths are caused by the metastasis of the cancer rather than the primary tumor itself. Yet, the underlying mechanisms of ...

Subcutaneous Angiotensin II Infusion using Osmotic Pumps Induces Aortic Aneurysms in Mice

Osmotic pumps continuously deliver compounds at a constant rate into small animals. This article introduces a standard protocol used to induce aortic aneurysms via subcutaneous infusion of angiotensin II (AngII) from implanted osmotic pumps. This protocol includes calculation of AngII amount and dissolution, osmotic pump filling, implantation of osmotic pumps subcutaneously, observation after pump implantation, and harvest of aortas to visualize aortic aneurysms in mice. Subcutaneous infusion of AngII through osmotic pumps following this protocol is a reliable and reproducible technique to induce both abdominal and thoracic aortic aneurysms in mice. Infusion durations range from a few days to several months based on the purpose of the study. AngII 1,000 ng/kg/min is sufficient to provide maximal effects on abdominal aortic aneurysmal formation in male hypercholesterolemic mouse models such as apolipoprotein E deficient or low-density lipoprotein receptor deficient mice. Incidence of abdominal aortic aneurysms induced by AngII infusion via osmotic pumps is 5 - 10 times lower in female hypercholesterolemic mice and also lower in both genders of normocholesterolemic mice. In contrast, AngII-induced thoracic aortic aneurysms in mice are not hypercholesterolemia or gender-dependent. Importantly, multiple features of this mouse model recapitulate those of human aortic aneurysms.

Subcutaneous implantation of osmotic pumps provides a convenient approach for prolonged and consistent delivery of compounds. This approach has been used extensively to study both abdominal and thoracic aortic aneurysms in mice.

Osmotic pumps continuously deliver compounds at a constant rate into small animals.  This article introduces a standard protocol used to induce aortic ...

Depletion of Mouse Cells from Human Tumor Xenografts Significantly Improves Downstream Analysis of Target Cells

The use of in vitro cell line models for cancer research has been a useful tool. However, it has been shown that these models fail to reliably mimic patient tumors in different assays1. Human tumor xenografts represent the gold standard with respect to tumor biology, drug discovery, and metastasis research 2-4. Tumor xenografts can be derived from different types of material like tumor cell lines, tumor tissue from primary patient tumors4 or serially transplanted tumors. When propagated in vivo, xenografted tissue is infiltrated and vascularized by cells of mouse origin. Multiple factors such as the tumor entity, the origin of xenografted material, growth rate and region of transplantation influence the composition and the amount of mouse cells present in tumor xenografts. However, even when these factors are kept constant, the degree of mouse cell contamination is highly variable.
Contaminating mouse cells significantly impair downstream analyses of human tumor xenografts. As mouse fibroblasts show high plating efficacies and proliferation rates, they tend to overgrow cultures of human tumor cells, especially slowly proliferating subpopulations. Mouse cell derived DNA, mRNA, and protein components can bias downstream gene expression analysis, next-generation sequencing, as well as proteome analysis 5. To overcome these limitations, we have developed a fast and easy method to isolate untouched human tumor cells from xenografted tumor tissue. This procedure is based on the comprehensive depletion of cells of mouse origin by combining automated tissue dissociation with the benchtop tissue dissociator and magnetic cell sorting. Here, we demonstrate that human target cells can be can be obtained with purities higher than 96% within less than 20 min independent of the tumor type.

Human tumor xenografts are vascularized and infiltrated by cells of mouse origin during the growth phase in vivo. To circumvent the bias caused by these contaminating cells during downstream analysis, we developed a method allowing for comprehensive depletion of all mouse cells by magnetic cell sorting.

The use of in vitro cell line models for cancer research has been a useful tool. However, it has been shown that these models fail to reliably mimic ...

Expression of Exogenous Cytokine in Patient-derived Xenografts via Injection with a Cytokine-transduced Stromal Cell Line

Patient-derived xenograft (PDX) mice are produced by transplanting human cells into immune deficient mice. These models are an important tool for studying the mechanisms of normal and malignant hematopoiesis and are the gold standard for identifying effective chemotherapies for many malignancies. PDX models are possible because many of the mouse cytokines also act on human cells. However, this is not the case for all cytokines, including many that are critical for studying normal and malignant hematopoiesis in human cells. Techniques that engineer mice to produce human cytokines (transgenic and knock-in models) require significant expense before the usefulness of the model has been demonstrated. Other techniques are labor intensive (injection of recombinant cytokine or lentivirus) and in some cases require high levels of technical expertise (hydrodynamic injection of DNA). This report describes a simple method for generating PDX mice that have exogenous human cytokine (TSLP, thymic stromal lymphopoietin) via weekly intraperitoneal injection of stroma that have been transduced to overexpress this cytokine. Use of this method provides an in vivo source of continuous cytokine production that achieves physiological levels of circulating human cytokine in the mouse. Plasma levels of human cytokine can be varied based on the number of stromal cells injected, and cytokine production can be initiated at any point in the experiment. This method also includes cytokine-negative control mice that are similarly produced, but through intraperitoneal injection of stroma transduced with a control vector. We have previously demonstrated that leukemia cells harvested from TSLP-expressing PDX, as compared to control PDX, exhibit a gene expression pattern more like the original patient sample. Together the cytokine-producing and cytokine-negative PDX mice produced by this method provide a model system that we have used successfully to study the role of TSLP in normal and malignant hematopoiesis.

Described here is a method for producing exogenous cytokine in patient-derived xenograft (PDX) mice via weekly intraperitoneal injection of a cytokine-transduced stromal cell line. This method broadens the utility of PDX and provides the option for transient or sustained exogenous cytokine delivery in a multitude of PDX models.

Patient-derived xenograft (PDX) mice are produced by transplanting human cells into immune deficient mice. These models are an important tool for studying ...