Abstract
Purpose:
To develop and characterize a porcine model of liver cancer, which could be used to test new locoregional therapies.
Methods:
Liver tumors were induced in 18 Oncopigs (transgenic pigs with Cre-inducible TP53R167H and KRASG12D mutations) by using an adenoviral vector encoding the Cre-recombinase gene. The resulting tumors (n=60) were characterized on multiphase contrast-enhanced CT, angiography, perfusion, micro CT, and necropsy. Transarterial embolization was performed using 40–120 micron (4 pigs) or 100–300 micron Embospheres (4 pigs). Response to embolization was evaluated on imaging. Complications were determined based on daily clinical evaluation, laboratory results, imaging, and necropsy.
Results:
Liver tumors developed at 60/70 (86%) inoculated sites. Mean tumor size was 2.1 cm at 1 week. Microscopically, all animals developed poorly differentiated to undifferentiated carcinomas, accompanied by a major inflammatory component, which resemble undifferentiated carcinomas of the human pancreatobiliary tract. Cytokeratin and vimentin expression confirmed epithelioid and mesenchymal differentiation respectively. Lymph node, lung, and peritoneal metastasis were seen in some cases. On multiphase CT, all tumors had a hypovascular center, and 17/60 (28%) had a hypervascular rim. After transarterial embolization, non-contrast CT showed retained contrast in the tumors. Follow up contrast-enhanced scan showed reduced size of tumors embolized using either 40–120 micron or 100–300 micron Embospheres, while untreated tumors showed continued growth.
Conclusion:
Liver tumors can be induced in a transgenic pig, and can be successfully treated using bland embolization.
Introduction
Current intra-arterial therapies for liver cancer or liver metastases — embolization, chemoembolization, and radioembolization — have several limitations, including a high rate of recurrence, damage to normal liver, and inability to treat extrahepatic disease. A wide range of new therapies have been proposed to address these limitations, including new embolic agents (1, 2), devices for local drug delivery (3–5), intra-arterial drugs (6–10), intra-arterial virus (11), and intravascular ablation modalities (12–15).
Realistic animal models are critical to develop and optimize innovative locoregional therapies for liver cancer. Existing animal models do not always accurately reflect outcomes in humans. Hepatocellular carcinoma (HCC) can be induced in rats and mice (16, 17). However, lobar embolization in rats (17) results in liver and lung necrosis, and a mortality rate of 56%. New cancer therapies developed in rodents have a high failure rate when translated to humans (18). The rabbit VX2 model is currently the most commonly used model for testing intra-arterial therapies in the liver. However, rabbits generally have different physiology (19, 20), and usually require higher weight-based drug dosing (21, 22) than humans.
This paper presents a new pig model of liver cancer, using the Oncopig (23, 24), a transgenic pig with Cre-recombinase-inducible TP53R167H and KRASG12D mutations. Unlike rodents and rabbits, pigs have similar size, physiology (19, 20), drug dosing (21, 22), and immune response (25, 26) compared to humans. Locoregional therapy in pigs can be performed using the same size catheters and devices that are used in humans. This porcine model could enable rapid testing and iteration of new devices, drugs, and techniques that are not yet ready for human trials.
Materials and Methods
Animals
The Institutional Animal Care and Use Committee approved all research procedures. The animal facility is AAALAC accredited and operates in compliance with the Guide for the Care and Use of Laboratory Animals (27). 18 female Oncopigs (transgenic pigs with Cre-inducible TP53R167H and KRASG12D mutations) were obtained from the University of Illinois, or the National Swine Resource and Research Center at the University of Missouri. All procedures and imaging were performed under general anesthesia, with peri-operative analgesia. Tumor induction was performed when the pigs were 8 – 22 weeks old. The overall study design is shown in Supplemental Figure 1.
Tumor induction (in situ method)
An 18 G core biopsy of the liver was obtained under CT or ultrasound guidance (1 or 2 cm core length, Temno Evolution, Merit Medical, South Jordan, UT), using co-axial technique (18 pigs). TP53R167H and KRASG12D expression was induced by incubating the core biopsy with an adenoviral vector carrying the Cre recombinase gene (109 pfu Ad5CMVCre-eGFP, University of Iowa Viral Vector Core) for 20 minutes at room temperature, in phosphate buffered saline containing 15 mM calcium chloride (total fluid volume of 1 ml). Gelatin sponge (Gelfoam, Pfizer) was then added using a 3-way stopcock, and the mixture (virus, core biopsy, gelatin) was injected percutaneously back into the pig’s liver through the biopsy needle, which was kept in place. At least two sites were inoculated in each liver. Inoculation sites were selected to be far apart as possible, easy and safe to access, and deep enough to avoid leakage of injected material into the peritoneum.
Tumor induction (cell line method)
Autologous hepatocyte-derived cell lines were created (8 pigs), as previously described (28, 29). Briefly, liver resection was performed, and hepatocytes were purified. The hepatocytes were transformed using Ad5CMVCre-eGFP, then passaged in cell culture. Transformed cells (6×107) were then mixed with gelatin sponge and injected percutaneously into the pig’s liver, under CT or ultrasound guidance. At least two sites were inoculated in each liver.
Multiphase contrast enhanced CT
Five-phase contrast-enhanced CT was performed 1 and 2 weeks after tumor inoculation to monitor tumor growth and response to therapy (18 pigs; as described below, a subset of pigs was treated using transarterial embolization immediately after the 1 week scan). Non-contrast CT of the abdomen and pelvis was obtained. Omnipaque 300 (2 ml/kg, max 150 ml) was power injected at 2–3 ml/sec. The early arterial phase CT scan was obtained when the abdominal aorta reached 150 Hounsfield units. The late arterial phase was obtained 15 seconds after the early arterial phase. The portal venous phase was obtained 25 seconds after the late arterial phase scan. The delayed phase scan was obtained 90 seconds after portal venous phase. All scans were obtained at 120 kVp.
Liver tumor perfusion
Hepatic artery and portal vein perfusion of the largest liver tumor in each pig (18 pigs) was estimated based on quantitative measurements from the multiphase CT, as previously described (30).
Angiography, cone beam CT, and micro CT
From femoral access, under fluoroscopic guidance, a 4 F catheter was advanced into the celiac artery, and an arteriogram was performed. The 4 F catheter or a 2.4 F microcatheter were advanced into the hepatic artery, and an arteriogram was performed. Cone beam CT arteriogram was obtained during a breath hold after administering a paralytic agent (rocuronium 1–1.6 mg/kg IV).
To obtain high resolution micro CT of tumor arteries, yellow silicone contrast (Microfil, Flow Tech Inc, Carver, MA) was injected into the hepatic artery, two weeks after tumor inoculation, in a single pig with untreated liver tumors. The liver tumor was resected, and ex vivo micro CT was performed at 85 µm resolution, using a microCAT II microCT (Imtek Corp, Oak Ridge, TN).
Transarterial embolization
One week after tumor inoculation, selective transarterial embolization of one liver tumor per pig was performed to stasis, using 40–120 µm (4 pigs) or 100–300 µm (4 pigs) Embospheres (Merit Medical, South Jordan, Utah). Follow up CT scan and necropsy were performed 1 week after embolization.
Safety evaluation
Complications were determined based on daily clinical evaluation post-treatment, CT scan, pre- treatment and pre-necropsy laboratory results (liver function tests, basic metabolic panel, complete blood count), and necropsy.
Pathology
After the 2-week post-inoculation scan, animals were euthanized, and liver tumors were macroscopically examined, harvested, and fixed in 10% neutral buffered formalin (18 pigs). Complete necropsies were performed in 2 pigs. Following formalin fixation, sections of tumor were processed into paraffin blocks, and sectioned at 5 micron thickness. Hematoxylin and eosin (H&E)-stained sections were reviewed by both human (OB, GA) and veterinary (AOM, SM) pathologists. Representative formalin-fixed paraffin-embedded tissue sections were immunolabeled with antibodies against cytokeratin AE1/AE3, vimentin, Iba1, arginase, and CD31 (Supplemental Table 1).
Statistical analysis
Tumor sizes were compared using two-tailed t-tests. Proportions were compared using Fisher’s exact test or a chi-squared test. Statistical analysis was performed in Excel 2016 and Mathematica 12. P values less than 0.05 were considered significant.
Results
Imaging
Liver tumors developed at 60 out of 70 sites (86%) that were inoculated. Mean tumor size was 2.1 cm (range: 0.3 – 4 cm) at 1 week. There was no significant difference in inoculation success rate (p=0.06) or tumor size (p=0.47) for sites inoculated using the in situ method (n=48) or the cell line method (n=22). There was a trend towards lower inoculation success rate with the cell line method (73%) compared to the in situ method (92%).
On multiphase contrast-enhanced CT, all tumors had a hypovascular center (Figure 1), and 17 of 60 (28%) had a hypervascular rim (Supplemental Figure 2A). Two tumors invaded the hepatic vein, and one tumor invaded the portal vein (Supplemental Figure 2B).
Figure 1.
Imaging characteristics of an Oncopig liver tumor (arrows). Arterial (A), portal venous (B), and delayed phase (C) contrast-enhanced CT images are shown, as well as celiac angiogram (D) and cone beam CT (E). The tumor is hypovascular on CT, but mildly hypervascular on angiography and cone beam CT.
Perfusion
Hepatic artery and portal vein perfusion to Oncopig liver tumors (Supplemental Figure 3) overlaps with previously collected perfusion data on human HCC (30) and colorectal cancer liver metastases (31). 35% of human HCCs, and 83% of human colorectal liver metastases fall within the range of hepatic artery and portal vein coefficients seen in Oncopig liver tumors.
Micro CT
Micro CT of untreated Oncopig liver tumors (Figure 2) shows that the tumors recruit blood supply from the hepatic artery, and develop new tumor vascularity. The scan resolution is 85 µm, which is smaller than many embolic particles.
Figure 2.
Micro CT of a liver tumor in the Oncopig, after intra-arterial injection of radiopaque silicone (maximum intensity projection image, 10 mm thick slab, with tumor vessels colorized red). The image shows a tumor feeding artery (white arrow; 0.3 mm; 3 other feeding vessels not shown), branching intratumoral arteries (black arrows), disorganized intratumoral vessels (white arrowheads), and tiny vascular lakes (black arrowheads). The intratumoral vessels measure up to 0.5 mm, and the vascular lakes measure up to 1 mm. Scan resolution is 85 µm, and a 1 cm bar and 200 µm circle are shown for scale.
Pathology
Grossly, the tumors were solid, pale tan lesions involving the liver. Histopathology revealed poorly differentiated to undifferentiated carcinomas, accompanied by a major inflammatory component, in all 18 pigs (Figure 3). H&E-stained sections (46 tumors) showed 4 different tumor morphologies: 1. Numerous atypical epithelioid cells organized in hypercellular anastomosing islands (15 of 46 tumors). 2. Small number of atypical epithelioid cells present as individual cells or very small clusters (21 of 46 tumors). 3. Numerous spindle cells organized in hypercellular bundles with a smaller epithelioid cell component (7 of 46 tumors). 4. Inflammation only, without atypical cells (3 of 46 tumors). There was no association between the tumor induction technique (in situ or cell line) and tumor morphology (p=0.44).
Figure 3.
Pathology of Oncopig tumors. (A) Gross pathology shows a pale tan, well circumscribed soft tissue nodule (arrow) in the liver (1 cm scale bar). (B) Hematoxylin and eosin stained section shows that the tumor is composed of solid islands of atypical cells with eosinophilic cytoplasm and large nuclei (50 μm scale bar). (C) Cytokeratin AE1/AE3 expression (brown) in atypical cells confirms epithelioid differentiation (50 μm scale bar). (D) Vimentin immunopositivity (brown) confirms mesenchymal differentiation (50 μm scale bar). (E) Lung metastasis (200 μm scale bar). (F) Lymph node metastasis (100 μm scale bar). Additional immunohistochemistry is shown in Supplemental Figure 4.
All tumors had a strong inflammatory response that was composed of lymphocytic and plasmacytic inflammatory infiltrates, often surrounding neoplastic cells and infiltrating portal regions of the liver. In contrast, necrotic portions of the tumor were surrounded by histiocytes and neutrophils. Multinucleated giant cells were often noted in the tumors.
Immunohistochemistry revealed that neoplastic cells with an atypical epithelioid morphology had two distinct immunophenotypes. They were either cytokeratin AE1/AE3 and vimentin immunopositive (Figure 3C and D) or AE1/AE3 immunonegative and vimentin immunopositive. The spindle cells were AE1/AE3 immunonegative and vimentin immunopositive (Supplemental Figure 4). Cytokeratin expression indicates epithelial differentiation, and vimentin expression indicates mesenchymal differentiation. Loss of cytokeratin expression, with maintenance of malignant cytologic features, suggests epithelial-to-mesenchymal transition. Tumor cells were immunonegative for Iba1, arginase, and CD31.
Metastases to lymph nodes, peritoneum, and lung were seen in some cases (Figure 3E and F). Two pigs had a complete necropsy, and an additional three pigs had incidentally noted extrahepatic tumors that were submitted to pathology. Histologically confirmed metastases were seen in the peritoneum (5 of 5 pigs), lymph nodes (2 of 5 pigs), lung (1 of 5 pigs), and pleura (1 of 5 pigs). It is important to note that this study was not designed to evaluate metastases, and most pigs did not have a complete necropsy, so the actual incidence of metastases is unknown. In addition, it is unknown whether these are true metastases. The peritoneal tumors could be due to leakage into the peritoneum during liver inoculation, and the lung tumor could be due to inoculation into the hepatic vein.
Overall features were similar to those of poorly differentiated to undifferentiated carcinomas, with or without multinucleated giant cells, of the human pancreatobiliary tract (32, 33).
Embolization
After transarterial embolization, non-contrast CT showed retained contrast in the tumors (Figure 4). Follow up contrast-enhanced scan showed (Figure 4) decreased size and enhancement of embolized tumors (Figure 4).
Figure 4. Transarterial embolization.
Contrast enhanced CT (A) shows an Oncopig liver tumor (pre- treatment). After transarterial embolization of the liver tumor using 40–120 µm Embospheres, immediate post-procedure non-contrast CT shows retained contrast in the tumor (B). One week later, follow up contrast-enhanced scan shows decreased size and enhancement of the liver tumor (C).
Untreated tumors grew by 0.3 cm between 1 and 2 weeks after inoculation (Figure 5). In the same time interval, tumors shrank by 0.2 cm after embolization using 40–120 µm Embospheres (p=0.03), and they shrank by 0.3 cm after embolization using 100–300 µm Embospheres (p=0.07).
Figure 5.
Change in liver tumor size, 1 week after transarterial embolization, compared to untreated tumors. After embolization using 40–120 µm Embospheres, tumor size decreased (p=0.03), compared to untreated tumors. Other pairwise comparisons were not statistically significant. The error bars show standard error of the mean.
Complications
The complication rate was 25% (2 of 8 pigs). One pig treated with 40–120 micron Embospheres developed post-embolization syndrome (vomiting) requiring euthanasia. One pig treated with 100– 300 micron Embospheres died during sedation for terminal imaging, and was found to have a large right pleural effusion, liver infarct, and progressive extrahepatic metastases (peritoneum, lung). Histopathology showed Embospheres in both the hepatic artery and portal vein, which could explain the liver infarct.
Discussion
This paper presents a promising new pig liver tumor model, with fast and reproducible site-specific tumor induction. The liver tumors are hypovascular, with a hypervascular rim, macrovascular invasion, and metastases in some cases. Histopathology showed poorly differentiated to undifferentiated carcinomas with a major inflammatory component. Oncopig liver tumor blood supply is similar to human liver tumors, and the tumors respond to transarterial embolization. The tumors can invade the portal vein or hepatic vein, like HCC. Comparison to other animal models is summarized in Supplemental Table 2.
Oncopig liver tumor vascularity was characterized on both perfusion CT and micro CT. Perfusion parameters in human liver tumors predict response to radioembolization of colorectal liver metastases (31), and survival after embolization of HCC (30). This paper shows that Oncopig liver tumors have hepatic artery and portal vein perfusion characteristics that overlap with both human colorectal liver metastases and HCC.
Micro CT of Oncopig liver tumors shows how the tumors recruit blood supply and develop tumor neovascularity. The tumors contain vascular lakes, which are also seen in human HCC (34), where they predict response to chemoembolization. Characterization of microscopic tumor vasculature down to the arteriole level (at the size of embolization particles) could provide insights into tumor angiogenesis and outcomes after therapy.
Necropsy and micro CT can provide detailed feedback on the causes of treatment failure and complications, which are not easily obtained in humans. For example, in an Oncopig with liver infarcts after embolization, necropsy showed embolic particles in both the hepatic artery and portal vein, suggesting an angiographically occult arterioportal fistula as the cause of the infarct.
A major inflammatory component was seen in all of the Oncopig liver tumors. Undifferentiated carcinomas in humans can also contain an inflammatory infiltrate (32, 33). Subcutaneous and intramuscular tumors in the Oncopig also contain a major inflammatory component, which is due to an antitumor T-cell response (35). Future experiments should address whether these inflammatory pig tumors could serve as a good model for the anti-tumor immune response in humans.
The cell(s) of origin of these neoplasms could not be determined. Expression of cytokeratins and vimentin indicate epithelial and mesenchymal differentiation, respectively, and co-expression of these markers, in association with tumor morphology observed on H&E, is most consistent with a primary undifferentiated carcinoma undergoing epithelial-to-mesenchymal transition. Alternatively, the heterogenous appearance of the tumors could be due to transformation of more than one cell type.
Major epithelial populations of the liver that can give rise to carcinomas are hepatocytes and biliary epithelial cells. Arginase, a hepatocyte marker, was not expressed by tumor cells, but this lack of expression does not rule out a hepatocellular origin, as expression can be lost in undifferentiated tumors. There was no difference in histologic appearance of tumors generated using the cell line technique (which involved transforming purified hepatocytes) and the in situ technique (which involved transforming a core biopsy of the liver, which contains both epithelial and mesenchymal cell types).
The Oncopig model has previously been used to generate subcutaneous and intramuscular tumors (28, 36), but not liver tumors. Other pig liver tumor models are available (37), but they require more than 1 year for tumor development, which makes it difficult to rapidly test and iterate new therapies.
The simplified single-step tumor inoculation method reported here (in situ method) is technically simpler to implement than the cell line method, which requires liver resection and cell culture, and the rabbit VX2 model, which requires growing tumors subcutaneously before transferring them to the liver. The in situ tumor inoculation method can be performed by any interventional radiologist with access to an animal lab.
The Oncopig liver tumor model can be used to develop new image-guided therapies, such as tumor- targeting intra-arterial drug carriers (38), and intra-arterial local immunotherapy (39). One limitation of the Oncopig model is that it is more expensive than small animal models (but possibly easier to translate to human trials). Another limitation is that the inflammatory, poorly differentiated, rapidly growing tumors might not be a good model for well differentiated or slowly growing tumors. However, the similarity in perfusion to human liver tumors, and the response to bland embolization, suggest that it will be a good model for arterially-directed therapies and local drug delivery. The robust antitumor immune response could be helpful for developing new immunotherapy techniques.
In conclusion, liver tumors can be induced in a transgenic pig, and can be successfully treated using bland embolization. The Oncopig liver tumor model is a potential alternative to the rabbit VX2 model, especially in cases where animal size or physiology are important.
Supplementary Material
Acknowledgements
This work was supported by the Office of the Assistant Secretary of Defense for Health Affairs, through the Peer Reviewed Cancer Research Program, under Award No. W81XWH-16–1-0338. Opinions, interpretations, conclusions and recommendations are those of the author and are not necessarily endorsed by the Department of Defense. Some pigs were supplied by the National Swine Resource and Research Center, which was supported by NIH grant U42 OD011140. Our animal facility was supported in part by GE, and core facilities were funded in part through an NIH/NCI Cancer Center Support Grant (P30 CA008748).
Govind Srimathveeravalli provided valuable advice. Megin Reilly, Lee-Ronn Paluch, Jacqueline Candelier, Brian Hanna, Stephanie Harris-Ash, Alvin Lopez, and Brian Cuevas helped with animal experiments. Liqun Cai helped with cell culture experiments. Afsar Barlas, Maria Jiao, and Mesruh Turkekul performed immunohistochemistry. Regina Schwind helped coordinate experiments.
Footnotes
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Disclosures
FEB is a co-founder of Claripacs, LLC. He received research support (investigator-initiated) from GE. He received research supplies (investigator-initiated) from Bayer, Steba Biotech, and Terumo. He received a research grant and speaker fees from Society of Interventional Oncology, which were sponsored by Guerbet. He attended research meetings sponsored by Guerbet. He is an investor in Labdoor, Qventus, CloudMedx, Notable Labs, and Xgenomes. He is the inventor and assignee on US patent 8233586, and is an inventor on US provisional patent applications 62/754,139 and 62/817,116.
SBS is a consultant for Johnson & Johnson, Aperture Medical, XACT Robotics, Innoblative, Endoways, and Varian. He received grants from GE Healthcare, AngioDynamics, Elesta, and Johnson & Johnson. He is a shareholder in Aspire Bariatrics, Aperture Medical, Johnson & Johnson, Immunomedics, Impulse, Motus GI, and Progenics.
RCG receives research support from Guerbet USA LLC and Janssen Research & Development LLC.
HY is an advisory board member of Genetech and BD Medical.
LBS has an equity consulting role in Sus Clinicals, Inc.
Contributor Information
Fuad Nurili, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
Sebastien Monette, Laboratory of Comparative Pathology, Memorial Sloan Kettering Cancer Center, The Rockefeller University, Weill Cornell Medicine, New York, NY.
Adam O. Michel, Laboratory of Comparative Pathology, Memorial Sloan Kettering Cancer Center, The Rockefeller University, Weill Cornell Medicine, New York, NY
Achiude Bendet, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
Olca Basturk, Department of Pathology, Memorial Sloan Kettering Cancer Center, New York, NY.
Gokce Askan, Department of Pathology, Memorial Sloan Kettering Cancer Center, New York, NY.
Christopher Cheleuitte-Nieves, Research Animal Resource Center, Sloan Kettering Institute, New York, NY.
Hooman Yarmohammadi, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
Aaron W. P. Maxwell, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
Etay Ziv, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
Kyle M. Schachtschneider, Department of Radiology, University of Illinois at Chicago, Chicago, Illinois.
Ron C. Gaba, Department of Radiology, University of Illinois at Chicago, Chicago, Illinois.
Lawrence B. Schook, Department of Animal Sciences, University of Illinois at Urbana-Champaign, Urbana, IL.
Stephen B. Solomon, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
F. Edward Boas, Interventional Radiology Service, Department of Radiology, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, NY 10065.
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