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Method Article

Portal Vein Injection of Colorectal Cancer Organoids to Study the Liver Metastasis Stroma

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DOI:

10.3791/62630

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September 3rd, 2021

 ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  ,  , 
1Adelaide Medical School, University of Adelaide, 2South Australian Health and Medical Research Institute (SAHMRI), 3Department of Pathology, Nagoya University Graduate School of Medicine, 4Division of Molecular Pathology, Center for Neurological Disease and Cancer, Nagoya University Graduate School of Medicine, 5Department of Gastroenterology, Graduate School of Medicine, The University of Tokyo, 6Translational and Clinical Research Institute, Newcastle University, 7International Center for Cell and Gene Therapy, Fujita Health University

In This Article

Summary

Portal vein injection of colorectal cancer (CRC) organoids generates stroma-rich liver metastasis. This mouse model of CRC hepatic metastasis represents a useful tool to study tumor-stroma interactions and develop novel stroma-directed therapeutics such as adeno-associated virus-mediated gene therapies.

Abstract

Hepatic metastasis of colorectal cancer (CRC) is a leading cause of cancer-related death. Cancer-associated fibroblasts (CAFs), a major component of the tumor microenvironment, play a crucial role in metastatic CRC progression and predict poor patient prognosis. However, there is a lack of satisfactory mouse models to study the crosstalk between metastatic cancer cells and CAFs. Here, we present a method to investigate how liver metastasis progression is regulated by the metastatic niche and possibly could be restrained by stroma-directed therapy. Portal vein injection of CRC organoids generated a desmoplastic reaction, which faithfully recapitulated the fibroblast-rich histology of human CRC liver metastases. This model was tissue-specific with a higher tumor burden in the liver when compared to an intra-splenic injection model, simplifying mouse survival analyses. By injecting luciferase-expressing tumor organoids, tumor growth kinetics could be monitored by in vivo imaging. Moreover, this preclinical model provides a useful platform to assess the efficacy of therapeutics targeting the tumor mesenchyme. We describe methods to examine whether adeno-associated virus-mediated delivery of a tumor-inhibiting stromal gene to hepatocytes could remodel the tumor microenvironment and improve mouse survival. This approach enables the development and assessment of novel therapeutic strategies to inhibit hepatic metastasis of CRC.

Introduction

Colorectal cancer (CRC) is a major cause of cancer mortality worldwide1. More than half of the CRC patients develop hepatic metastasis that occurs through the portal vein dissemination1. Currently, there are no effective therapeutics that can cure advanced liver metastasis, and most patients succumb to metastatic disease.

The metastatic niche or tumor microenvironment plays a key role in engraftment and growth of disseminated CRC cells2. Cancer-associated fibroblasts (CAFs), a prominent component of the tumor microenvironment, promote or restrain cancer progression through secreting growth factors, remodeling the extracellular matrix (ECM), and modulating immune landscapes and angiogenesis3,4,5. CAFs also confer resistance to chemotherapies and immunotherapies3. Moreover, CAFs regulate initiation and progression of CRC liver metastasis and predict prognosis in patients with CRC3,6,7,8. Thus, CAF-related factors could be exploited for the development of therapeutic strategies to inhibit CRC liver metastasis. However, the lack of satisfactory mouse models to study the metastatic tumor stroma has been a major obstacle to developing stroma-targeted therapies.

Currently, animal models to study CRC liver metastasis include primary CRC models that spontaneously develop hepatic metastasis and cancer cell transplantation models into the liver. Primary CRC mouse models, such as genetically engineered mouse models and colonic injection of cancer cells, rarely show metastasis to the liver9,10,11,12. Moreover, even if a liver metastasis is observed, these models show long latency from the primary tumor induction to metastasis, and potentially die of primary tumor burden12. To efficiently generate CRC liver metastases, cultured CRC cells are transplanted into the liver using three injection approaches: intra-splenic injection, direct intra-parenchymal injection into the liver, and portal vein injection. Intra-splenically injected cancer cells spread into the splenic vein, the portal vein, and ultimately to the liver13,14. However, the intra-splenic injection yields a lower tumor take ratio compared with other transplantation models15,16. With intra-splenic injection, surgical removal of the spleen is performed to avoid cancer growth in the spleen, which can potentially compromise immune cell maturation17. Furthermore, intra-splenic injection can also result in unintended tumor growth in the spleen and abdominal cavity18, complicating liver metastasis analyses. Direct intra-parenchymal injection into the liver efficiently induces hepatic metastasis16,19,20. Nonetheless, this approach does not fully recapitulate a biological step of liver metastasis that naturally occurs through portal vein dissemination. Using direct injection into the liver, entry of cancer cells into a non-portal, but systemic circulation can also result in multiple large lung metastases16. Although a majority of patients with CRC liver metastasis show multiple tumor nodules in the liver21, direct injection into a specific liver lobe generates a single tumor mass19,20. Portal vein injection or mesenteric vein injection, though technically challenging, allows efficient delivery of tumor cells into the liver in a manner that recapitulates the growth patterns seen in patients17. This strategy can minimize the possibility of secondary-site metastases and enables rapid growth of cancer cells in the liver, simplifying mouse survival analyses.

Historically, colorectal cancer cell lines such as mouse MC-38, human HT-29, and SW-620 were used to generate mouse models of hepatic metastasis22,23. However, these colorectal cancer cell lines do not induce a desmoplastic stromal reaction. Low stromal content in the tumors makes it difficult to investigate the biological roles of cancer-associated fibroblasts. Recent advances in CRC organoids and their transplantation have offered useful platforms to assess vital roles of the stroma in cancer progression24. Liver transplantation of CRC organoids generates a fibroblast-rich tumor microenvironment and has provided novel insights into stromal research6,25. Currently, portal or mesenteric vein injection of organoids has become a gold standard approach to generate CRC liver metastasis6,25,26,27,28. Nonetheless, to our knowledge, no previous papers have described detailed methods for the portal vein injection of colorectal tumoroids. Here, we present a methodology for using portal vein injection of CRC organoids to develop novel adeno-associated virus (AAV)-mediated stroma-directed therapy.

Hepatocytes are an important constituent of the metastatic tumor microenvironment in the liver and play a critical role in metastatic cancer progression29. Inspired by the success of AAV gene therapy approaches to induce protein expression in hepatocytes in non-neoplastic patients30,31, we investigated a similar approach but aimed at modifying the liver tumor microenvironment in CRC25. As such, we also describe herein the tail vein injection of AAV8 to induce expression of anti-tumorigenic proteins to modify the liver tumor microenvironment. The AAV8 serotype, designated by the choice of viral capsid protein during virus production, leads to high transduction efficiency specifically of hepatocytes (i.e., targeted gene expression in the liver tumor microenvironment)32. We have previously shown that Islr (immunoglobulin superfamily containing leucine-rich repeat) is a CAF-specific gene that induces bone morphogenetic protein (BMP) signaling, reduces CRC tumoroid growth, and promotes Lgr5+ intestinal stem cell differentiation25. We tested whether AAV8-mediated overexpression of the cancer-restraining stromal gene, Islr, in hepatocytes could attenuate hepatic metastasis progression by performing portal vein injection of CRC tumoroids in AAV8-Islr-treated mice.

In this paper, we first describe the tail vein injection procedure of liver tropic AAV. Then, we describe a method for tumoroid cell preparation and portal vein injection into the AAV-treated mice. Finally, we present approaches to monitor metastatic tumor progression to assess the efficacy of stroma-directed therapeutics.

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Protocol

All animal procedures in this article were reviewed and approved by the South Australian Health and Medical Research Institute Animal Ethics Committee (Approval number, SAM322).

1. Tail vein injection of adeno-associated virus

NOTE: Adeno-associated virus (AAV) should be handled as a biohazard under Biosafety Level 1 guidelines. Please refer to the published protocol for AAV preparation, purification, and titration33. Hepatocyte-tropic AAV, AAV834, encoding the cytomegalovirus (CMV) promoter-Islr gene, was used in this study25. To induce AAV-mediated overexpression, AAV dosing might require optimization depending on the promoter activity, gene, and mouse weight.

  1. Dilute AAV vector into 150 µL aliquots containing 1.0 x 1011 viral genomes, using sterile phosphate-buffered saline (PBS) and keep it on ice. Personal protective equipment should be worn to handle the AAV.
    NOTE: Repeated freeze and thaw cycle decreases virus titer and should be avoided. The stock viral solution should be stored in a -80 °C freezer.
  2. Turn on a heat box (animal warming chamber) to preheat to 35 °C.
  3. Hold a mouse and apply a topical anesthesia cream to the whole length of the tail at least 15 min prior to the tail vein injection.
    NOTE: This is an optional step and might not be necessary if it is not required by the animal ethics committee of the institute. Follow the protocol approved by the local animal ethics committee. In this experiment25, a Rosa26-Cas9 mouse was used for AAV injection and subsequent portal vein injection of CRC organoids. Given that tumoroids used in this study were derived from a Rosa26-Cas9 mouse (C57BL/6 x 129 genetic background), this mouse strain was also used as immunocompetent, syngeneic recipients of the tumor transplant. Male and female Rosa26-Cas9 mice (6- to 24-week-old) were used.
  4. Place the mouse into the heat box. Leave the mouse for up to 15 min to warm and dilate the tail veins.
  5. Gently secure the mouse in a rodent restrainer. Place the tail under a heat lamp to ensure full dilation of tail veins.
  6. Draw up 150 µL of diluted AAV (prepared in step 1.1) into a low dead space sterile syringe with a 27-30 G needle.
  7. Move the heat lamp and identify the lateral tail vein located on the sides of the tail. Put slight tension on the tail with the fingers so that the tail becomes straight.
  8. Slowly insert 2-3 mm of the needle, bevel up, into the vein. The needle should be almost parallel to the tail (up to 15° from the tail).
    NOTE: Blood influx into the syringe might be observed if the needle is successfully located into the vein.
  9. Inject slowly. If a resistance is felt or skin swelling is observed, remove the needle and re-insert above the first site or to the other lateral vein.
  10. Wait for about 5 s after the completion of the injection, and then slowly remove the needle. Immediately apply gentle pressure to the injection site with a clean gauze or paper towel until bleeding stops.
  11. Gently release the mouse into its cage. Monitor the animal to ensure bleeding has stopped.
    NOTE: Gene overexpression in the liver can be assessed 1 to 2 weeks after AAV tail vein injection. This can be confirmed, for example, by RNA in situ hybridization (ISH), immunohistochemistry (IHC), western blotting, or quantitative real-time polymerase chain reaction (qRT-PCR; Figure 1A,B). In a previously published study25, AAV8-Islr was used to overexpress mouse Islr gene in hepatocytes and the overexpression was detected by RNA ISH (Figure 1A,B).

2. Cell preparation for colorectal cancer organoids

NOTE: CRC organoids used for this experiment solely contain epithelial cells. Culture and generation of CRC organoids has been previously described25,35. In short, normal colonic epithelial cells were isolated from the colon of a Rosa26-Cas9 mouse using a crypt isolation buffer (5 mM EDTA (ethylenediaminetetraacetate) in ice-cold PBS), and then embedded in basement membrane matrix medium, and cultured in organoid growth medium as described in reference35. Then, Apc and Trp53 mutations were introduced to the colonic epithelial cells by overexpressing single-guide RNAs that target Apc and Trp53 using lentivirus expression protocol. Single organoid clones were handpicked25. ApcΔ/Δand Trp53Δ/Δ colon cancer organoids (AP tumoroids), were injected as 5.0 x 105 single cells in 100 µL of PBS with 10 µM Y-27632 into the portal vein per mouse, with organoid culture and single-cell preparation described below.

  1. Culture the CRC organoids in the basement membrane matrix medium domes in a 24-well plate or 10 cm dish 3-5 days before the portal vein injection to obtain 50-400 µm diameter organoids.
  2. Prepare sufficient amount of cell detachment solution by adding Y-27632 to 10 µM concentration, enough to digest the number of organoids being cultured for the injection. Pre-warm in a 37 °C water bath.
    NOTE: The cell detachment solution used in this protocol is a recombinant cell-dissociation enzyme mix and is used as a substitute for trypsin in organoid culture (see Table of Materials). It reduces cellular damage caused by cell dissociation compared to trypsin36. Y-27632 is a Rho kinase inhibitor and inhibits dissociation-induced cell death, thereby increasing single-cell survival37.
    1. For injection into up to five mice, prepare two tubes containing 40 mL of the cell detachment solution to digest 10-24 x 50 µL of basement membrane matrix domes with each dome containing approximately 300 organoids (50-400 µm diameter), i.e., each mouse is injected with 2-4.8 domes of organoids (equivalent to about 600-1440 organoids). The number of organoids necessary to obtain sufficient cell number is dependent on the organoid line, and thus should be optimized.
      NOTE: The second 40 mL tube enables a repeat digestion with fresh cell detachment solution to obtain dissociated single cells.
  3. Carefully aspirate the organoid medium from each well.
  4. Add 1 mL of ice-cold PBS to each well in a 24-well plate. If a 10 cm dish is used, add 10 mL of ice-cold PBS to the dish.
  5. Scratch off the basement membrane matrix medium using a P1000 pipette tip. Transfer the PBS/medium slurry to a 15 mL centrifuge tube.
  6. Rinse each well with the same amount of PBS to collect basement membrane matrix fragments and add to the 15 mL tube(s).
  7. Incubate for 5 min on ice. This incubation helps dissolve the basement membrane matrix medium.
  8. Centrifuge the tube at 400 x g for 5 min at 4 °C.
  9. Aspirate the supernatant ensuring not to disturb the pelleted medium and cells.
  10. Add 5 mL of pre-warmed cell detachment solution prepared in step 2.2 to the pellet, resuspend the pellet 10 times, and transfer it back into a 50 mL tube containing fresh, pre-warmed cell detachment solution.
  11. Place the tube in the 37 °C water bath. Incubate for 5 min.
  12. Centrifuge the tube at 400 x g for 3 min at 4 °C.
  13. Repeat steps 2.9-2.11.
    NOTE: Repeated enzymatic digestion allows for the efficient cell dissociation into single cells.
  14. Check whether the organoids are dissociated into single cells by pipetting 100 µL into a 96-well plate and observing under a microscope. If many organoids show cell clumps consisting of more than four cells, longer incubation with the cell detachment solution and physical dissociation by trituration with a 10 mL pipette may be necessary.
  15. Once most cells are single cells, add 4 mL of fetal bovine serum (FBS) into the 40 mL cell suspension to stop the digestion. Rinse a 40 µm cell strainer with 5 mL PBS.
  16. Pass the cell suspension through the cell strainer into a 50 mL collection tube to remove any cell clumps.
  17. Centrifuge the tube at 400 x g for 5 min at 4 °C.
  18. Aspirate the supernatant. Add 10 mL of cold PBS to the cell pellet and transfer it to a 15 mL centrifuge tube.
  19. Centrifuge the tube at 400 x g for 5 min at 4 °C.
  20. Aspirate the supernatant. Resuspend the pellet in 500 µL cold PBS with 10 µM Y-27632. Count the cells.
  21. Adjust the cell concentration to 5.0 x 105 single cells/100 µL, using PBS with 10 µM Y-27632. Place the tube on ice until portal vein injection is performed.
    ​NOTE: It is recommended to inject dissociated cells within 4 h of dissociation.

3. Portal vein injection of CRC organoids

NOTE: All surgical instruments and surgical gauzes must be autoclaved or sterilized before surgery. This protocol is modified from a previous protocol17. In this experiment25, portal vein injection was performed using Rosa26-Cas9 mice treated with AAV-mRuby2 or AAV-Islr in step 1.

  1. Prepare an aseptic surgical area using sterile drapes on a heating pad.
  2. Prepare surgical instruments (scissors and forceps), surgical and hemostatic sponge, 4-0 polyglactin suture, cotton buds, skin staplers, stapler applicator, saline, buprenorphine, and 33 G needle attached to a Hamilton syringe. Cut the hemostatic sponge into 1.0 cm x 1.0 cm pieces.
  3. Adjust the position of the light source to illuminate the surgical area.
  4. Inject 0.1 mg/kg body weight of buprenorphine subcutaneously to a mouse for surgical pain management.
  5. Anesthetize the mouse with isoflurane in an anesthesia chamber. Isoflurane concentration for induction and maintenance are usually 5% and 2.5%, respectively. Check for the absence of a reaction to toe pinch before commencing the next procedure.
  6. Shave the middle to the upper abdomen of the mouse with an electric shaver. Shaving should be performed in an area distant from the aseptic surgical area to avoid hair contaminating the site.
  7. Clean the incision site alternating with Betadine and 80% ethanol to sterilize the surgical area. Repeat three times.
  8. Place the mouse on a heating pad in a supine position with maintenance isoflurane anesthesia. Place a surgical drape with a hole over the abdomen of the mouse.
  9. Lift the abdominal skin with forceps and make a 2-3 cm skin incision at the midline with scissors, cutting the skin only (not underlying peritoneum). The incision should range from the mid-abdomen to the xiphoid process of the sternum. The incision should not go above the lower end of the xiphoid process.
  10. Fully lift the peritoneal wall with forceps and make a similar 2-3 cm incision to the peritoneum with scissors. Avoid cutting the intestine and the diaphragm.
  11. Soak the surgical gauze with warm saline and place it on the left side of the incision (on the left side of the mouse's body; to the surgeon's right).
  12. Gently pull the internal organs (small and large intestines) out using a cotton bud soaked with saline. Place the intestines on the gauze soaked with saline.
    NOTE: The mouse's left-side intestines (i.e., intestines on the surgeon's right) should be pulled out first, and then the mouse's right-side intestines (i.e., intestines on the surgeon's left) can be pulled out.
  13. Adjust the position of the intestines to visualize the portal vein. Cover the intestines with additional wet gauze to keep the intestines moist.
  14. Gently pull the intestine to the left side with the wet gauze and apply gentle tension to the left. This facilitates visualization of the portal vein (Figure 2A).
    NOTE: If visualization of the portal vein is difficult, gently adjusting the position of the stomach with a wet cotton bud might help the visualization.
  15. Gently pipette tumoroid suspension several times to obtain a homogenous cell suspension. Slowly draw up 100 µL of the cell suspension into a Hamilton syringe attached to a 33 G needle. Avoid air bubbles.
  16. Slowly insert the needle, bevel up, into the portal vein. The insertion depth along the needle should be 3-4 mm, with the needle angle almost parallel to the portal vein.
    NOTE: Injection should be performed into the well-visualized part of the portal vein (usually up to 2 cm away from the hepatic hilum). Avoid movement of the needle after it is fully inserted into the portal vein.
  17. Inject tumor cells for 30 s. The injection should be slowly performed to prevent occlusion of the portal vein. If the injection is successful, the color of the liver temporarily changes from red to white.
  18. Remove the needle slowly. Immediately apply gentle pressure to the injection site with a dry cotton bud and wait for 5 min.
  19. Remove the cotton bud and simultaneously apply a hemostatic sponge to the injection site. Hold hemostatic sponge with a cotton bud or forceps and apply gentle pressure for 5 more min.
  20. Remove the pressure to the hemostatic sponge and confirm that there is no bleeding from the injection site.
    NOTE: The bio-absorbable hemostatic sponge does not need to be removed. Trying to remove the gauze could cause re-bleeding from the injection site.
  21. If bleeding occurs, immediately perform pressure hemostasis with a cotton bud for about 10 min. Then, apply an additional hemostatic sponge for a further 5 min.
    NOTE: If uncontrollable blood loss is observed, the mouse should be euthanized according to the protocol approved by the animal ethics committee of the institute.
  22. Remove the surgical gauzes on the intestines. Using a syringe filled with 5 mL saline, squirt saline to the intestines to prevent organ adhesion.
    NOTE: Do not apply saline to the portal vein injection site. This could cause re-bleeding.
  23. Gently place the intestines back inside the abdominal cavity.
  24. Suture the peritoneum using 4-0 polyglactin sutures.
  25. Lift up both sides of the skin with forceps. Apply skin staplers to close the skin incision. Be careful not to staple the intestine.
  26. Turn off the isoflurane but keep the oxygen flow running. Carefully monitor the mouse. When the mouse awakens, place the mouse in an empty cage on a heating pad. The mouse typically awakens within 5 min.
  27. Carefully monitor the mouse until it is conscious and ambulating normally.
  28. Subcutaneously inject 0.1 mg/kg buprenorphine to the mouse 4 h after the surgery.
  29. Inject 0.1 mg/kg buprenorphine to the mouse every 24 hours for the subsequent 2 days.
  30. Carefully monitor the mice daily for a week after surgery. Check sutures and wound healing.
  31. days after the surgery, remove skin staplers using a stapler remover.

4. Assessment of tumor growth kinetics by in vivo bioluminescent imaging

NOTE: If Firefly-expressing tumoroids are used for injection, metastatic tumor progression can be monitored weekly by in vivo imaging as described38,39. Luciferase expressed by cancer cells could elicit immune responses against the cancer cells and limit tumor growth40. Thus, caution is warranted in analyzing immune phenotypes and cancer progression in a mouse model using luciferase-expressing tumor cells.

  1. Prepare a 30 mg/mL solution of D-luciferin using sterile PBS. Protect it from light. D-luciferin should be stored in aliquots at -20 °C until use.
  2. Shave the abdomen and thorax with an electronic shaver. This can be done up to 1 day before in vivo imaging.
  3. Inject 150 mg/kg body weight of D-luciferin intraperitoneally into mice (i.e., if the mouse body weight is 30 g, inject 150 µL of the D-luciferin solution).
  4. Place the mice in an anesthesia chamber and anesthetize them. Use 5% of isoflurane for induction and 2%-3% for maintenance.
  5. min later, place the mice in a lateral position (right side up). Acquire bioluminescence images using an in vivo imaging system (IVIS) as described previously38,39.
    NOTE: Mice are more stable in a lateral position compared to the supine position. Therefore, a lateral position is preferable to obtain a luminescence image from a consistent focal plane.
  6. Place the mice in an empty cage and monitor their recovery.
  7. Define regions of interest on the upper abdomen using Living Image Software as described38. Quantify total flux as a surrogate for tumor cell number.

5. Survival analysis and tissue collection

  1. Monitor mice carefully for clinical symptoms of metastases such as a distended abdomen.
  2. Euthanize a mouse by CO2 inhalation once it reaches humane endpoints.
    NOTE: Use a study endpoint approved by the animal ethics committee of the institute. To determine a humane endpoint, a clinical record sheet was used; scores were calculated by one point being given for the presence of each of the following observations: weight loss > 15%, hunched posture, ruffled coat, dehydration, decreased movement, distended abdomen, or facial grimace. Once a score of 3 was reached, the mice were humanely euthanized.
  3. Immediately following euthanasia, perform transcardial perfusion fixation with 30-50 mL of 10% formalin in a fume hood, as described41. Make several small incisions (up to 1 cm each) with scissors into the normal part of the liver before perfusion to generate outlets for blood and formalin. Upon perfusion, the liver color will change from red to brown.
    NOTE: If micrometastases in macroscopically normal liver areas are a subject of study, we recommend researchers to not cut the liver, but to snip the superior vena cava instead. However, when we have used this method, fixation of the liver seems poorer when compared with cutting the liver. Especially if RNA in situ hybridization (ISH) is planned to be performed on liver sections, we recommend making incisions into the liver before intra-cardiac injection of formalin, as described above. This enables sufficient fixation of the liver tissue, resulting in preservation of RNA integrity for ISH.
  4. Place the liver and lung tissues into 10% formalin and fixate overnight. Replace the formalin with 70% ethanol, followed by paraffin-embedding.
  5. Perform hematoxylin and eosin staining to histologically evaluate tumor area. Perform immunohistochemistry for stromal markers of interest. Perform Picro-Sirus Red staining to evaluate collagen-positive areas.
    NOTE: The ImageJ software42 can be used to quantify immunohistochemistry and Picro-Sirius Res staining data. A color deconvolution function and the MRI fibrosis tool can be used to evaluate 3,3'-Diaminobenzidine (DAB)-positive areas and Picro-Sirius red-positive areas, respectively.

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Results

To induce AAV-mediated overexpression of a tumor-restraining stromal gene, Islr4,25,43,44, in hepatocytes, we intravenously injected Islr-encoding AAV8. 1.0 x 1011 viral genomes (vg) of AAV8-Islr, or as a control, AAV8-mRuby2, was injected into the adult mouse tail vein (Figure 1A). Two weeks after the tail vein inject...

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Discussion

In this study, we have shown that portal vein injection of mouse CRC organoids reproducibly generates fibroblast-rich liver metastases that mimic histological features of human CRC hepatic metastases. Furthermore, when combined with stroma-directed therapeutics such as AAV8-mediated gene therapy, this preclinical model serves as a useful tool to assess therapeutic effects on mouse survival and tumor growth.

There are, at least, two critical steps in the protocol. Firstly, it is important to pr...

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Disclosures

The authors declare no conflicts of interest.

Acknowledgements

This study was supported by grants from the National Health and Medical Research Council (APP1156391 to D.L.W., S.L.W.) (APP1081852 to D.L.W., APP1140236 to S.L.W., APP1099283 to D.L.W.,); Cancer Council SA Beat Cancer Project on behalf of its donors and the State Government of South Australia through the Department of Health (MCF0418 to S.L.W., D.L.W.); a Grant-in-Aid for Scientific Research (B) (20H03467 to M.T.) commissioned by the Ministry of Education, Culture, Sports, Science and Technology of Japan; AMED-CREST (Japan Agency for Medical Research and Development, Core Research for Evolutional Science and Technology (19gm0810007h0104 and 19gm1210008s0101 to A.E.); the Project for Cancer Research and Therapeutic Evolution (P-CREATE) from AMED (19cm0106332h0002 to A.E.); Japan Society for the Promotion of Science Overseas Challenge Program for Young Researchers (to H.K.), Takeda Science Foundation Fellowship (to H.K.), Greaton International Ph.D. Scholarship (to H.K.), Lions Medical Research Foundation Scholarship (to K.G.).

We thank Dr. Leszek Lisowski at Vector and Genome Engineering Facility (VGEF), Children's Medical Research Institute (CMRI) (NSW, AUSTRALIA) for producing recombinant AAV vectors.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
10% FormalinSigmaHT501128
15 mL centrifuge tubeCorning430791
33-gauge needleTSKLDS-33013For portal vein injection
4-0 vicryl sutureETHICONJ494G
40-µm cell strainerCorning431750
5 mL SyringeBD302130Used to apply saline to the intestine after portal vein injection
50 mL centrifuge tubeCorning430829
50 mL syringeTERUMOSS*50LELuer lock syringe for perfusion fixation
70% Isopropyl alcohol wipeBriemar5730
Anaesthesia machineDarvall9356
αSMA antibodyDAKOM0851Clone 1A4. 1/500 dilution for immunohistochemistry
BuprenorphineTROYN/Ailium Temvet Injection, 300 µg/ml Buprenorphine
Cotton budsJohnson & JohnsonN/AJohnson's pure cotton bud applicators. Need to be autoclaved before use.
D-luciferinBiosynthL-8220
Electric shaverSold by multiple suppliers
ForcepsSold by multiple suppliers
Hamilton syringeHAMILTON81020For portal vein injection
Heat box (animal warming chamber)DatesandMK3
Heat lampSold by multiple suppliers
Hemostatic spongePfizer09-0891-04-015Gelfoam absorbable gelatin sponge, USP, 12-7 mm
India inkTalens44727000
Injection syringe and needleBD326769For tail vein injection
Islr probe (RNAscope)ACD450041
IsofluraneHenry Schein988-3244
IVIS Spectrum In Vivo Imaging SystemPerkin Elmer124262
Living Image SoftwarePerkin Elmer128113
MatrigelCorning356231
MRI fibrosis toolN/AN/Ahttps://github.com/MontpellierRessourcesImagerie/imagej_macros_and_scripts/wiki/MRI_Fibrosis_Tool
Phosphate-buffered saline (PBS)SigmaD8537
RNAscope kitACD322300
Rodent restrainerSold by multiple suppliers
Rosa26-Cas9 mouseThe Jackson Laboratory024858
SalinePfizerPHA19042010
ScissorsSold by multiple suppliers
Skin staplersAble ScientificAS590289 mm wound clips
Stapler applicatorAble ScientificAS590269 mm wound clip applicator
Stapler removerAble ScientificAS59037Wound clip remover
Surgical drapeMultigate29-220
Surgical gauzeSentry MedicalGS001
Topical anesthesia creamEMLAN/AEMLA 5% cream, 25 mg/g lignocaine and 25 mg/g prilocaine
TrypLE ExpressGibco12605028Recombinant cell-dissociation enzyme mix
Y-27632Tocris1254

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