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Cancer Research

Non-invasive PET/MR Imaging in an Orthotopic Mouse Model of Hepatocellular Carcinoma

Published: August 31, 2022 doi: 10.3791/63958
* These authors contributed equally


Here, we present a protocol to create orthotopic hepatocellular carcinoma xenografts with and without hepatic artery ligation and perform non-invasive positron emission tomography (PET) imaging of tumor hypoxia using [18F]Fluoromisonidazole ([18F]FMISO) and [18F]Fluorodeoxyglucose ([18F]FDG).


Preclinical experimental models of hepatocellular carcinoma (HCC) that recapitulate human disease represent an important tool to study tumorigenesis and evaluate novel therapeutic approaches. Non-invasive whole-body imaging using positron emission tomography (PET) provides critical insights into the in vivo characteristics of tissues at the molecular level in real-time. We present here a protocol for orthotopic HCC xenograft creation with and without hepatic artery ligation (HAL) to induce tumor hypoxia and the assessment of their tumor metabolism in vivo using [18F]Fluoromisonidazole ([18F]FMISO) and [18F]Fluorodeoxyglucose ([18F]FDG) PET/magnetic resonance (MR) imaging. Tumor hypoxia could be readily visualized using the hypoxia marker [18F]FMISO, and it was found that the [18F]FMISO uptake was higher in HCC mice that underwent HAL than in the non-HAL group, whereas [18F]FDG could not distinguish tumor hypoxia between the two groups. HAL tumors also displayed a higher level of hypoxia-inducible factor (HIF)-1α expression in response to hypoxia. Quantification of HAL tumors showed a 2.3-fold increase in [18F]FMISO uptake based on the standardized value uptake (SUV) approach.


Hepatocellular carcinoma (HCC) is the sixth most diagnosed cancer and the third most common cause of death from cancer worldwide, with more than 900,000 new cases and 800,000 deaths in 20201. The major risk factor is cirrhosis, which occurs as a result of viral infections (hepatitis B and C viruses), alcohol abuse, diabetes, and non-alcoholic steatohepatitis2. The management of HCC is rather complex, and several treatment options are available, including surgical resection, thermal or chemical ablation, transplantation, transarterial chemoembolization, radiation, and chemotherapy, depending on the disease staging2,3. HCC is a chemotherapy-refractory tumor with disease recurrence in up to 70% of patients following curative-intent therapy2.

Despite the high degree of tumor heterogeneity, HCC is associated with two common outcomes: (i) HCC is very hypoxic, and (ii) tumor hypoxia is linked to greater tumor aggressiveness and treatment failure. The uncontrolled proliferation of HCC cells results in a high oxygen consumption rate that precedes vascularization, thus creating a hypoxic microenvironment. Low intra-tumoral oxygen levels then trigger a range of biological responses that influence tumor aggressiveness and treatment response. Hypoxia-inducible factors (HIFs) are often recognized as the essential transcriptional regulators in the response to hypoxia2,3. Hence, the ability to detect hypoxia is crucial to visualize neoplastic tissues and identify the inaccessible sites, which require invasive procedures. It also helps to better understand the molecular changes that lead to tumor aggressiveness and improve patient treatment outcomes.

Molecular imaging using positron emission tomography (PET) is commonly used in the diagnosis and staging of many cancers, including HCC. In particular, the combined use of dual-tracer PET imaging involving [18F]Fluorodeoxyglucose ([18F]FDG) and [11C]Acetate can significantly increase overall sensitivity in HCC diagnosis4,5. Imaging of hypoxia, on the other hand, can be achieved by using the commonly used hypoxic marker [18F]Fluoromisonidazole ([18F]FMISO). In clinical practice, the non-invasive assessment of hypoxia is important to differentiate between various types of tumors and regions for radiation therapy planning6.

Preclinical imaging has become an indispensable tool for the non-invasive and longitudinal evaluation of mouse models for different diseases. A robust and highly reproducible HCC model represents an important platform for preclinical and translational research into the pathophysiology of human HCC and the assessment of novel therapies. Together with PET imaging, in vivo behaviors can be elucidated to provide important insights at the molecular level for any given timepoint. Here, we describe a protocol for the generation of hepatic artery ligation (HAL) orthotopic HCC xenografts and analysis of their in vivo tumor metabolism using [18F]FMISO and [18F]FDG PET/MR. The incorporation of HAL makes a suitable model of transgenic or chemically induced HCC mice xenografts to study tumor hypoxia in vivo, as HAL can effectively block the arterial blood supply to induce intra-tumoral hypoxia7,8. In addition, unlike ex vivo immunohistochemical staining using pimonidazole, changes in tumor metabolism as a result of hypoxia can be readily visualized and accurately quantified non-invasively using PET imaging, enabling longitudinal assessment of treatment response or gauging of the emergence of resistance3,7,8. Our method shown here allows the creation of a robust hypoxic HCC model together with non-invasive monitoring of tumor hypoxia using PET/MR imaging to study HCC biology in vivo.

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All animal studies were carried out in accordance with the Committee on the Use of Live Animals in Teaching and Research (CULATR) in the Centre for Comparative Medicine Research (CCMR) at the University of Hong Kong, a program accredited by the Association for the Assessment and Accreditation of Laboratory Animal Care International (AAALAC). The animals used in the study were female BALB/cAnN-nu (Nude) mice at the age of 6-8 weeks, weighted at 20 g ± 2 g. Food and water were provided ad libitum.

1. Subcutaneous injection of human hepatocellular carcinoma cell lines

NOTE: MHCC97 is a human HCC cell line that is used to generate the subcutaneous HCC xenograft model. MHCC97L cells are obtained from the Liver Cancer Institute, Zhongshan Hospital of Fudan University, Shanghai, the People's Republic of China9 and authenticated by short tandem repeat (STR) profiling.

  1. Culture MHCC97L cells and maintain in culture media supplemented with 10% fetal bovine serum (FBS) and 1% penicillin and streptomycin in a T-75 cell culture flask at 37 °C in a humidified incubator supplied with 5% CO2 (see Table of Materials). Change the culture media every 2 days and harvest the cells at 80%-90 % confluency.
    NOTE: The starting cell number is 1.5 x 106 cells, and the cells are used within 6-8 passages.
  2. Store the culture media and trypsin-ethylenediaminetetraacetic acid (trypsin-EDTA) at 4 °C in the dark until use. Pre-warm the culture media to 37 °C and the trypsin-EDTA to room temperature before use. Refer to the Table of Materials for all reagents and consumables used in this section.
  3. Remove the culture media. Rinse the cell layer with 10 mL of sterile phosphate-buffered saline (DPBS, see Table of Materials).
  4. Add 5 mL of trypsin-EDTA to the culture flask and incubate the cells at 37 °C. Check the cells under an inverted microscope at 20x magnification (see Table of Materials) until the cell layer is dispersed and completely detached from the flask. Make sure that the cells are not left in trypsin-EDTA for more than 15 min.
  5. Add 5 mL of culture media to the culture flask. Aspirate the cells by pipetting gently. Transfer the cell suspension to a centrifuge tube and spin at 1,107 x g for 5 min (see Table of Materials).
  6. Remove the supernatant and gently resuspend the cell pellet with 1 mL of PBS. Pipette thoroughly to obtain a single-cell suspension. Determine the cell number using the automated cell counter (see Table of Materials) and prepare the cell suspension to a concentration of 5 x 106 cells/mL in PBS.
  7. Draw up 200 μL of cell suspension with a 25 G needle in a 1 mL syringe. Each syringe contains 1 x 106 MHCC97L cells (in 200 μL). Keep the syringe on wet ice.
  8. Anesthetize the mouse with an intraperitoneal (i.p.) injection of a mixture of ketamine (87.5 mg/kg) and xylazine (12.5 mg/kg). Ensure proper anesthesia using the toe-pinch reflex. Make sure to apply eye lubricant to the mouse to prevent drying and the potential formation of corneal ulcers. Place the mouse in a supine position on a sterile pad (see Table of Materials).
  9. Sterilize the dorsal region near the lower flank (either left or right side) of the mouse with 70% ethanol (see Table of Materials). Lift the skin up with forceps and gently insert the needle bevel up underneath the skin.
  10. Advance the needle 4-5 mm along the subcutaneous plane from the puncture site to avoid accidental leakage of the cell suspension from the injection site upon withdrawal of the needle. Make sure that the needle insertion is not too deep under the skin, which may cause accidental organ damage.
  11. Release the forceps and carefully and slowly discharge the contents of the syringe, taking care not to penetrate the opposite side. Pull out the needle to check for a swell on the injection. Place the mouse back into the cage sitting on top of a pre-warmed heating pad to assist with recovery from the anesthesia and to regain mobility. Continue to monitor the mouse until the mouse is fully awake and maintains sternal recumbency.
  12. Monitor the tumor growth and wellbeing of the mouse on a weekly basis. Measure the tumor volume (V) using a caliper (see Table of Materials). Calculate and record the tumor volume using the following formula10:
    V = (W2× L)/2
    where W is the width and L is the length of the tumor.
  13. Between 4-6 weeks post injection, check that a significant size of subcutaneous tumor, between 800-1000 mm3, is obtained. Make sure that the total tumor load does not exceed 10% of the body weight.

2. Orthotopic liver implantation and hepatic artery ligation

  1. Prepare the following surgical reagents and tools in a disinfected biological safety cabinet: 10 mL of culture media in a dish, 10% povidone-iodine solution, 70% ethanol, autoclaved surgical instruments, i.e., sharp scissors, spring scissors, forceps (curved fine and straight blunt), needle holder, scalpel blade, 6-0 and 5-0 nylon suture, a heating pad.
  2. Autoclave all the surgical instruments and handle them only on a sterile bench. Provide the adequate and necessary pre-, intra-, and post-operative care to the mice throughout the surgical procedures (see steps below).
    1. For pre-operative pain relief, inject the mouse with a subcutaneous injection of buprenorphine (0.1 mg/kg) at least 30 min prior to the start of the surgical procedure. For post-operative care and management, inject the mouse with a subcutaneous injection of buprenorphine (0.1 mg/kg) for 3 days continuously, once every 8-12 h. Refer to the Table of Materials for all reagents and consumables used in this section.
  3. Euthanize the subcutaneous tumor-bearing mouse with an i.p. injection of pentobarbital (330 mg/kg). Make an incision on the skin of the tumor site using a scalpel blade. Excise the subcutaneous tumor block and transfer it immediately to the culture media. Perform this step as quickly as possible.
  4. Cut the tumor block into 1 mm3 small tumor cubes using sharp surgical scissors. Keep all the tumor cubes in the culture media. Avoid using the core region of the subcutaneous tumor block.
  5. Anesthetize the mouse that will be undergoing the subsequent orthotopic tumor implantation surgery with an i.p. injection of a mixture of ketamine (87.5 mg/kg) and xylazine (12.5 mg/kg). Make sure that at least 30 min has elapsed after giving the buprenorphine prior to anesthetizing the mouse. Make sure to apply eye lubricant to the mouse to avoid drying and the potential formation of corneal ulcers.
  6. Place the mouse in a supine position on a sterile pad that is on top of a pre-warmed heating pad. Monitor and record the physiological state of the mouse every 15 min until the end of the surgical procedure.
  7. Extend the limbs of the mouse. Secure them with tape to maximize exposure of the ventral abdomen and thorax. Sterilize the entire abdominal skin of the mouse with a 10% povidone-iodine solution, followed by 70% ethanol. Assess the anesthetic depth of the mouse by checking for no response to toe pinching-the pedal withdrawal reflex.
  8. Perform a laparotomy by making an approximately 10 mm midline incision to access the peritoneum using sharp scissors. Clamp and pull the xiphoid process toward the head with forceps and apply the subcostal retractors to open the surgical field. Make sure that a proper incision is made to enable a good surgical view of the operation site.
  9. Retract the median and left lobes upward with a wet gauze swab. Move the intestines outside the abdomen onto the left side of the mouse and cover them with a wet gauze swab. Perform hepatic artery ligation (HAL) on the common hepatic artery (CHA), which originates from the pancreatic head to its root in the hepatic pedicle, under a magnifying lamp for optimized visualization.
  10. Retract the median and left lobes back/downward with a wet gauze swab to expose the left lobe. Make an incision of roughly 2 mm in length and depth on the surface of the left lobe with a sterile scalpel blade.
    NOTE: It is also possible to use the median lobe of the liver for tumor implantation to minimize accidental tears or laceration by trauma to neighboring abdominal organs.
  11. Immediately insert a tumor fragment into the liver incision with sterile forceps. Bury the tumor so that it sits securely in the liver. Apply a figure-of-eight suture with a 6-0 nylon suture over the incision site to ensure proper tumor fragment implantation into the liver and hemostasis. Perform this operation as quickly as possible to avoid excessive bleeding. Upon completion, close the abdomen incision with interrupted 5-0 sutures.
  12. Place the mouse back in the cage sitting on top of a pre-warmed heating pad to assist with recovery from anesthesia and to regain the righting reflex. Continue to monitor and record the physiological state of the mouse every 15 min until the mouse is fully awake and maintains sternal recumbency.
  13. Repeat the procedure until all mice have gone through orthotopic tumor implantation. Provide post-operative care to all mice by continuously monitoring their physiological state daily for the next 3 days after the surgical operation. Give buprenorphine (0.1 mg/kg) every 8-12 h.

3. Setting up of PET/MR calibrations and workflow

NOTE: Imaging is performed using a preclinical PET/MR 3T system (see Table of Materials).

  1. Use 5% isoflurane (1 L/min medical O2) to anesthetize the mouse in an induction chamber. For each scan, carefully place the anesthetized mouse on an imaging bed with continuous heating; first, scan with MR (scout view) as an anatomical reference for static [18F]FMISO or [18F]FDG PET scans, followed by anatomic reference MR imaging.
  2. Create a sequential PET/MR imaging workflow in the software (see Table of Materials) to include a static PET scan, T1-weighted (attenuation correction) and T2-weighted (anatomic reference) MR imaging, and PET reconstruction 1 day before the scheduled imaging session.
  3. To acquire PET, select 400-600 keV level discrimination, F-18 study isotope, 1-5 coincidence mode, and 20 min scans.
  4. To acquire whole-body T1-weighted MR, use gradient echo-3D with TE = 4.3 ms, TR = 16 ms, field of view (FOV) = 90 mm x 60 mm, number of excitations (NEX) = 3, 28 slices with 0.9 mm thickness, and voxel size = 0.375 mm3 x 0.375 mm3 x 0.9 mm3.
  5. To acquire whole-body T2-weighted MR, use fast-spin echo 2D with TE = 71.8 ms, TR = 3000 ms, FOV = 90 mm x 60 mm, NEX = 5, 28 slices with 0.9 mm thickness, voxel size = 0.265 mm3 x 0.268 mm3 x 0.9 mm3.
  6. To reconstruct PET, select Tera-Tomo (TT3D) algorithm, iterations = 8, subsets = 6, 1-3 coincidence mode, and with decay, dead-time, random, attenuation, and scatter corrections to create images with an overall voxel size of 0.3 mm3.
  7. Perform a PET activity test before the start of the experiment to check the accuracy of PET quantitation11.
    1. Fill a 5 mL syringe with 5-8 MBq [18F]FMISO diluted in water or saline as per the manufacturer's recommendation. Use a dose calibrator to record the activity of the syringe (see Table of Materials). Note the time of measurement.
    2. Draw a volume of interest (VOI) on the reconstructed image to cover the whole syringe. Compare the recovered activity from the image to the value obtained from the dose calibrator. Proceed with imaging studies when the recovered activity is accurate within ±5%.

4. [18F]FMISO and [18F]FDG injection

  1. Obtain a clinical dose of [18F]FMISO from the supplier 30 min before the start of the imaging studies. Wear the appropriate personal protective equipment (PPE), such as a lab coat, gloves, over spectacles, a film badge, and ring dosimeters when handling radioactive materials. Change gloves frequently to avoid cross-contamination of the radioactive substances.
  2. Place the radioactive stock solutions in the lead storage container behind an L-block table-top shield. Dispense an aliquot of [18F]FMISO and dilute with sterilized saline. Ensure that the total activity concentration is 18-20 MBq in 100 μL.
  3. Draw up the [18F]FMISO with a 1 mL syringe with needle (see Table of Materials). Record the radioactivity and time shown on the dose calibrator.
  4. Record the mouse weight before the injection of the radiotracer. Use 5% isoflurane (1 L/min medical O2) to anesthetize the mouse, then carefully inject the prepared [18F]FMISO via the tail vein. Record the injection time and residue activity of the syringe to account for decay correction. Place the mouse back in the cage for 180 min to allow [18F]FMISO uptake prior to the PET scan. Allow the mouse to recover for 1 day after [18F]FMISO PET.
    NOTE: Despite the very different in vivo pharmacokinetic profiles for [18F]FMISO and [18F]FDG, both radiotracers are radiolabeled with F-18, which is a PET radionuclide with a half-life of 109.77 min (< 2 h). Since the PET signal originates from F-18, half of the initial injected radioactivity will be lost every 2 h. In this case, the residual signal 24 h post injection can no longer be detected by the PET scanner. In addition, the 1 day gap between both [18F]FMISO (day 1) and [18F]FDG (day 3) injections will allow complete decay of [18F]FMISO when [18F]FDG PET is performed, hence no overlapping of [18F]FMISO signal in the [18F]FDG PET scan.
  5. Repeat steps 4.1.-4.4. for the [18F]FDG injection on day 3, with the exception that a total activity concentration of 6-8 MBq in 100 μL is injected with a 60 min uptake before the start of the PET scan.
  6. Calculate the injected [18F]FMISO or [18F]FDG activity using the following formula11:
    Injected activity (MBq) = Activity in the syringe before injection - Activity in the syringe after injection

5. PET/MR acquisition

  1. Turn on the air heater to warm the mouse imaging bed before the scheduled PET scan. Use 5% isoflurane (1 L/min medical O2) to anesthetize the mouse in an induction chamber.
  2. Once properly anesthetized and checked using the toe-pinch response, carefully and immediately transfer the mouse to the heating bed and maintain anesthesia at 2%-2.5% isoflurane. Place the mouse head-prone onto the bite bar and apply eye lubricant to both the eyes to avoid drying and the potential formation of corneal ulcers. Cover the imaging bed with a plastic sheet to maintain the bed's surrounding temperature.
  3. Monitor and record the respiratory rate and body temperature of the mouse using the in-house respiratory pad and thermal probe, respectively. Ensure that the mouse's body temperature is maintained at 37 °C while the respiratory rate is kept in the range of 40-50 breaths per min (bpm) by adjusting the isoflurane level.
  4. Perform a scout view to locate the mouse position inside the scanner. Adjust the mouse bed position to cover the whole mouse body, with the torso region located at the center FOV of the MR.
  5. To start a PET scan, select PET Acquisition in the study list window, then choose Scan Range on Previous Acquisition to use the pre-determined mouse bed position from the scout view. Click on Prepare > OK to automatically move the mouse bed from MR to PET and start the acquisition.
  6. Under the Radiopharmaceutical Editor, select the appropriate radionuclide, then enter the details of the injection dose and time before and after [18F]FMISO or [18F]FDG injection, as measured in step 4.3. and step 4.4. Enter the weight of the mouse under the Subject Information menu.
  7. Proceed with MR acquisitions upon completion of the PET scan. To do so, select Prepare to move the mouse to MR from PET. Click OK to continue.
  8. Once all the scans are completed, select Home to move the imaging bed back to the default position.
  9. Turn off the anesthesia and flush the imaging bed with medical O2. Check the mouse pedal withdrawal reflex. Transfer the mouse from the imaging bed back to a clean housing cage placed on top of a heating pad to assist with recovery from anesthesia and to regain the righting reflex.
  10. Under the Raw Scan, select PET Acquisition to load the acquired raw PET data for PET reconstruction. Select MM for use as a material map. Proceed with PET reconstruction using the parameter as described in step 3.6.
  11. Strictly adhere to the local and institutional guidelines when handling mice after PET imaging studies. Treat all used consumables, e.g., syringes, needles, gloves, wipes, and biological waste, e.g., cage bedding and fecal matter, that have been in direct contact with the mouse as radioactive waste, and handle with care according to the local regulations.

6. PET image analysis

  1. Open the Image Analysis software (see Table of Materials), and select Load DICOM Data to open the PET and MR images.
  2. Drag the corresponding PET and MR images to the software display window, and select Automatic Registration to perform automatic co-registration of the resulting PET and MR images.
    1. Under the Registration Setup menu, select Rigid transformation. Use Shift and Rotation in the Rigid/Affine menu.
    2. Under the Global Role Selection menu, click on MM as Reference and Static PET Scan as Reslice.
    3. Inspect the co-registered PET/MR images to ensure proper alignment. If necessary, use Manual Registration to adjust the images manually.
    4. Locate the orthotopic tumor in the liver using the MR image. Use Interpolated Ellipse ROI to draw a VOI on the tumor, using the MR image for reference. Transfer the created tumor VOI from the MR to the PET image. Select the Brush Tool and Eraser Tool to carefully define the VOI border slide-by-slide. Make sure to avoid spillover of radioactivity uptake from the liver on the PET image.
    5. Use Ellipsoid VOI to create a 3 mm3 VOI on the liver as a reference organ. Make sure to avoid areas of visible hepatic vascular and biliary structures by using the MR image as a guide.
    6. Select Show ROI Table to rename all the drawn VOIs. Record all the necessary data, e.g., radioactivity (kBq/mL) and tumor volume (mm3), in a spreadsheet. Save a copy of the VOI drawings and archive both the raw and reconstructed imaging data to an external hard drive when completed.
    7. Calculate the standardized uptake value (SUV) for all VOIs using the following equation11:
      SUV = CPET(t)/(ID/BW)
      where CPET(t) is the measured activity in the VOI, ID is the injected dose measured in kBq, and BW is the mouse body weight in kg, assuming a tissue density of 1 g/mL.

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Representative Results

To obtain a suitable tumor block for successive orthotopic implantation, stable clones were first generated by subcutaneous injection of 200 μL of cell suspension in DPBS (containing MHCC97L cells) into the lower flank of nude mice (Figure 1A). Tumor growth was monitored and, when tumor size reached 800-1000 mm3 (around 4 weeks post injection), mice were euthanized, and the resulting tumor block was cut into approximately 1 mm3 fragments for subsequent liver orthotopic implantation into another batch of nude mice (n = 6). Mice were randomized into two groups: control (C1, n = 3) and hepatic artery ligation (H2, n = 3). HAL was performed by tying a fine thread around the main branch of the hepatic artery. C1 mice were spared from HAL prior to orthotopic implantation. This manipulation led to tumor hypoxia in H2 but not C1 mice, and the tumor hypoxic status could be monitored non-invasively using the PET hypoxic tracer [18F]FMISO. PET/MR studies showed that an increase in tumor [18F]FMISO uptake was observed only in the H2 mice, whereas, using the glycolytic marker [18F]FDG, tumor uptake remained similar between the two groups (Figure 1B).

HAL-induced hypoxia in tumors was further validated by probing the expression levels of HIF-1α12, and comparisons were made between the groups. Consistent with a more hypoxic tumor after HAL, the H2 group exhibited higher HIF-1α expression than the C1 group (optical density: 0.17 vs. 0.13, H2 vs. C1, respectively), which corroborates with their tumor [18F]FMISO uptake (Figure 2A). [18F]FMISO uptake was quantified using the SUV-based approach. H2 tumors showed a 2.3-fold increase in [18F]FMISO uptake when compared to C1 tumors (SUVmax: 1.2 vs. 2.8, respectively, Figure 2B). Similarly, HAL also resulted in a higher liver SUV uptake in H2 than C1 mice (Figure 2C). Taken together, we show here that HAL can effectively induce tumor hypoxia in HCC orthotopic xenografts, and tumor hypoxia can be non-invasively monitored using [18F]FMISO PET imaging, supported by the ex vivo immunohistochemistry marker HIF-1α expression. In addition, the incorporation of MR imaging offers an excellent soft tissue contrast to enable clear delineation of the tumor from the liver, making accurate PET quantification possible.

Figure 1
Figure 1: In vivo PET/MR imaging of orthotopic HCC mice xenografts. (A) Schematic of subcutaneous and orthotopic xenograft creations and PET imaging studies in orthotopic MHCC97L tumors. (B) Representative maximum intensity projection (MIP) PET images of [18F]FMISO and [18F]FDG in mice bearing orthotopic MHCC97L tumors (C1) without or (H2) with HAL. Blue circles indicate the location of the tumor. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Analysis of mice bearing orthotopic MHCC97L tumors with and without HAL. (A) Representative co-registered [18F]FMISO PET/MR images, immunohistochemistry staining for HIF-1α, and hematoxylin and eosin (HE) in tumor sections. (B-C) Quantitative analysis of [18F]FMISO uptake in the tumor and liver. N = 3 for each group. Values of SUV are presented as mean ± standard error of the mean (SEM). Please click here to view a larger version of this figure.

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In this study, we described the procedures to perform HAL on liver orthotopic HCC xenografts using subcutaneous tumors, along with methods for the non-invasive monitoring of tumor hypoxia in orthotopic xenografts using [18F]FMISO and [18F]FDG PET/MR. Our interest lies in the metabolic imaging of various cancer and disease models for early diagnosis and treatment response evaluation11,13,14,15. To date, the creation of HAL HCC xenografts and their in vivo tumor metabolism have rarely been described in the literature, which prompted us to investigate the metabolic characteristics of these tumor models using PET imaging.

The successful establishment of orthotopic xenografts with HAL as a robust mice model to study hypoxia in HCC represents an important aspect for studying HCC biology in vivo. Hypoxia is known to stimulate cancer malignancy. Moreover, intra-tumoral hypoxia has been associated with enhanced proliferation, metastasis, and radio- and chemoresistance and warrants thorough characterization of the response to hypoxic conditions7. While subcutaneous xenografts are often used to study HCC tumorigenesis or treatment strategies, orthotopic models can better recapitulate human HCC development since they reflect more accurately the tumor microenvironment, particularly the influences on vascularization and tumor-stroma interactions toward HCC metastasis12. The incorporation of HAL into HCC orthotopic mice allows the induction of intra-tumoral hypoxia by blocking the hepatic arterial bloody supply to the tumor7. Such animal models enable mechanisms underlying the effects of HAL on HCC to be elucidated and create new avenues for effective therapeutics targeting the hypoxia pathway in HCC. For orthotopic implantation, we utilized the tumor cube from the subcutaneous tumor instead of direct injection of HCC cell suspensions. We have found that direct cell injections often cause a small volume of leakage from the injection site, even though the procedure was carried out as slowly as possible with a low injection volume. This may potentially lead to peritoneal metastasis, which is detrimental to animal wellbeing, ultimately affecting the overall study outcome. On the other hand, implantation of a small tumor block would overcome the issue, although care needs to be taken to ensure that the tumor is securely sutured after implantation. When isolating small tumor cubes from the subcutaneous tumor block, it is also advisable to avoid the core regions of the solid tumor, which are relatively less vascularized and more metabolically stressed due to hypoxia and nutrient deprivation. Doing so will reduce the inclusion of dead tumor cells that appear to be necrotic within the small tumor cube and will maintain more consistent tumor growth rates across individual mice.

Hypoxia is a prognostic factor of cancer resistance, and monitoring changes in hypoxia using PET allows the detection and quantification of hypoxic tissues in high sensitivity and specificity. An important consideration for [18F]FMISO and [18F]FDG PET involves the radiotracer uptake time and route of administration in mice. Both radiotracers have different in vivo pharmacokinetics, one being the physiological uptake difference, observed in the intestine/bladder and heart/bladder for [18F]FMISO and [18F]FDG, respectively, and the other being that [18F]FMISO accumulates in the hypoxic region of the tumor, and [18F]FDG is preferentially taken up in the high metabolic tumor region16. The high lipophilicity and slow hepatic clearance of [18F]FMISO require a 2-4 h post-injection time to obtain a good tumor-to-liver contrast17. We found that 3 h post injection is sufficient to differentiate and delineate tumor [18F]FMISO uptake from the liver for both hypoxic (H2) and control (C1) groups. In the case of [18F]FDG PET, a 1 h post-injection time is adequate to yield good contrast, as previously described13,15, and was employed in this study. Nevertheless, keeping a consistent radiotracer uptake time is imperative to ensure the reliability of PET results, especially for SUV-based analyses. Here, we used intravenous techniques to administer radiotracer to the mice. Successful tail vein injection is indicated by a visible blood flashback before the radiotracer infusion, where the needle is accurately positioned within the vein. A major drawback of this method is that the expected blood flashback may be difficult to observe due to hypotension, with variability observed between mice. Nevertheless, such difficulty can be overcome by warming the tail for a short period of 1-2 min prior to needle insertion, either with a warm washcloth or with a heat lamp to increase the blood flow and improve the vein visibility for successful injection.

With the increasing use of PET imaging to quantify radiotracer biodistribution in small animals, an important consideration to note is the use of an accurate and well-calibrated PET scanner to yield good, reproducible, and quantifiable PET data. An accurate PET system allows imaging studies to be performed in a more time-efficient and cost-effective manner and enables the implementation of the 3Rs principles (Replacement, Reduction, and Refinement) in animal research. For that reason, routine quality control inspections must be performed, preferably on a weekly basis, to examine the PET and MR components of the imaging system as per the manufacturer's recommendation. In particular, the accuracy for PET activity should be checked and recorded frequently, as well as before the start of an imaging experiment, to ensure the reliability of PET quantification. This is imperative for any longitudinal studies involving the quantitative evaluation of the tumor and tissues over an examination period to produce meaningful and comparable results. Calibration of the system is required when the PET activity accuracy is found to be out of the recommended range, as well as when misalignment of PET and MR images is discovered.

Although the techniques described here enable PET imaging of the hypoxic HCC xenografts for a wide range of in vivo investigations, some limitations should be considered when contemplating the use of these protocols. The creation of HAL HCC xenografts is a complex intra-abdominal surgical procedure to perform on 6-8-week-old immunodeficient mice. In addition, the diminutive hepatic artery in these young mice can be tough to locate, making the ligation process technically challenging. These procedures require suitably trained personnel to improve the surgery succession rate, as well as the survival of mice over a period of time, i.e., 4-6 weeks before xenografts form, while the provision of extensive animal care is expected throughout the surgery period, which are both time- and resource-intensive. It is also anticipated that the creation of orthotopic HCC xenografts using this method will require around 8 weeks prior to successful PET imaging, since implantation of the tumor cube is used rather than direct injection of cell suspensions to avoid peritoneal metastasis. Nevertheless, these limitations can be overcome with adequate planning and training of research staff. Also, the sample size of the experimental groups reported here are small, which might not be suitable for statistical analysis. However, our observations subjectively reveal a trend where HAL-induced hypoxia was measured in both the tumor and the liver in the H2 compared to the C1 group, which is supported by more prominent HIF-1α staining in the H2 tumor samples. We are currently working on expanding the tumor models with other human HCC cell lines. Also, therapeutic studies targeting the hypoxia pathway in HCC involving these xenografts are underway.

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The authors have no conflicts of interest to disclose.


We acknowledge the support of the Hong Kong Anticancer Trust Fund, Hong Kong Research Grants Council Collaborative Research Fund (CRF C7018-14E) for the small animal imaging experiments. We also thank the support of the Molecular Imaging and Medical Cyclotron Center (MIMCC) at The University of Hong Kong for the provision of [18F]FMISO and [18F]FDG.


Name Company Catalog Number Comments
0.9% sterile saline BBraun N/A 0.9% sodium chloride intravenous infusion, 500 mL
10# Scalpel blade RWD Life Science Co.,ltd S31010-01 Animal surgery tool
10% povidone-iodine solution Banitore 6.425.678 For disinfection
25G needle with a 1 mL syringe BD PrecisionGlide N/A 1 mL syringe with 25G needle for cell suspensions injections
5 mL syringe Terumo SS05L 5 mL syringe Luer Lock
70% Ethanol Merck 1.07017 For disinfection
Automated Cell Counter Invitrogen AMQAF2000 For automated cell counting
Buprenorphine HealthDirect N/A Subcutaneous injection (0.05-0.2 mg/kg/12 hours) for analgesic after surgery
Cell Culture Dish (60 mm diameter) Thermo Scientific 150462 For tumor tissue processing
Centrifuge Sigma 3-16KL, fixed-angle rotor 12311 For cell suspensions collection
Centrifuge Conical Tube Eppendorf EP0030122151 For cell suspensions collection
Culture media (Dulbecco’s modified Eagle’s medium) Gibco 10566024 high glucose, GlutaMAX™ Supplement
Digital Caliper RS PRO 841-2518 For subcutaneous tumor size measurement
Direct heat CO2 incubator Techcomp Limited NU5841 For cell culture
Dose calibrator Biodex  N/A Atomlab 500
DPBS (Dulbecco’s phosphate-buffered saline) Gibco 14287072 For cell wash and injection
Eye lubricant Alcon Duratears  N/A Sterile ocular lubricant ointment, 3.5 g
Fetal bovine serum (FBS) Gibco A4766801 Used for a broad range of cell types, especially sensitive cell lines
Forceps (curved fine and straight blunt) RWD Life Science Co.,ltd F12012-10 & F12011-13 Animal surgery tool
Heating pad ALA Scientific Instruments N/A Heat pad for mice during surgery
Insulin syringe Terumo 10ME2913 1 mL insulin syringe with needle for radiotracer injections
InterView fusion software Mediso Version 3.03 Post-processing and image analysis software
Inverted microscope Yu Lung Scientific Co., Ltd BM-209G For cells morphology visualization
Isoflurane Chanelle Pharma  N/A Iso-Vet, inhalation anesthetic, 250 mL
Ketamine Alfasan International B.V. HK-37715 Ketamine 10% injection solution, 10 mL 
Medical oxygen Linde HKO 101-HR compressed gas, 99.5% purity
nanoScan PET/MR Scanner Mediso  N/A 3 Tesla MR
Needle holder RWD Life Science Co.,ltd F31026-12 Animal surgery tool
Nucline nanoScan software Mediso Version 3.0 Scanner operating software
Nylon Suture (6/0 and 5/0) Healthy Medical Company Ltd 000524 & 000526 Animal surgery tool
Penicillin- Streptomycin Gibco 15140122 Culture media for a final concentration of 50 to 100 I.U./mL penicillin and 50 to 100 µg/mL streptomycin.
Pentabarbital AlfaMedic 13003 Intraperitoneal injection (330 mg/kg) to induce cessation of breathing of mice
Sharp scissors RWD Life Science Co.,ltd S14014-10 Animal surgery tool
Spring Scissors RWD Life Science Co.,ltd S11005-09 Animal surgery tool
Trypan Blue Solution, 0,4% Gibco 15250061 For cell counting
Trypsin-ethylenediaminetetraacetic acid (EDTA, 0.25%), phenol red. Gibco 25200072 For cell digestion
Xylazine Alfasan International B.V. HK-56179 Xylazine 2% injection solution, 30 mL



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Tan, K. V., Yang, X., Chan, C. Y., Shi, J., Chang, H. C., Chiu, K. W. H., Man, K. Non-invasive PET/MR Imaging in an Orthotopic Mouse Model of Hepatocellular Carcinoma. J. Vis. Exp. (186), e63958, doi:10.3791/63958 (2022).More

Tan, K. V., Yang, X., Chan, C. Y., Shi, J., Chang, H. C., Chiu, K. W. H., Man, K. Non-invasive PET/MR Imaging in an Orthotopic Mouse Model of Hepatocellular Carcinoma. J. Vis. Exp. (186), e63958, doi:10.3791/63958 (2022).

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