This work is supported by the SFB 824 subprojects C6 and C7 (DFG Sonderforschungsbereich 824), German Research Foundation, Bonn, Germany.

Note: The following steps have been performed in accordance with the guidelines of our institution and the local government of Bavaria, Germany. All attempts to reproduce this protocol must be made in adherence with the local guidelines for the humane treatment of animals, as dictated by local animal care and use committee. For example, many institutes require that sterile gloves are worn during rodent surgery.&#160;Furthermore, work with viruses must be performed according to local regulations, taking care for personal and environmental safety. Care must be taken to properly dispose of contaminated waste and to clean instruments and workspace (i.e., by autoclaving and cleaning with an appropriate antiviral disinfectant according to the manufacturer&#39;s instructions).
1. Preparations Before Beginning Surgery
<ol>
	<li>Sterilize surgical instruments (i.e., by autoclaving).</li>
	<li>Dilute the virus to the appropriate concentration, which will vary depending on the toxicity of the specific virus being used and the susceptibility of the rat strain. 
	Note: For example, when using vesicular stomatitis virus in male Buffalo rats, we typically inject 107 pfu in a 1 ml volume, which is the maximum tolerated dose in that model.
	<ol>
		<li>Prefill the virus (using aseptic technique) &#160;in 1 ml syringes, and attach 30 G needles, taking care to eliminate air from the syringe and needle.</li>
	</ol>
	</li>
	<li>Prepare the surgical space with all materials and equipment required, and spray the area with ethanol to disinfect. Fill a small receptacle with sterile saline solution (0.9% NaCl) for moistening gauze swabs and cotton-tipped applicators.</li>
</ol>
2. Preparation of the Rat
<ol>
	<li>Administer an analgesic (as specified by the local animal care and use committee), at the appropriate dose 30 min prior to beginning the surgical procedure. For example, administer a high dose of metamizole (100-200 mg/kg).</li>
	<li>Anesthetize the rat using inhalation (isoflurane) or injectable (i.e., a mixture of Medetomidine (0.15 mg/kg), Midazolam (2 mg/kg), and Fentanyl (0.005 mg/kg), also known as MMF) anesthesia. Administer isolflurane through a vaporizer at 1-3% and adjust as necessary to maintain normal respiration and heart rate.
	<ol>
		<li>Before proceeding, ensure that the rat is fully sedated and unable to feel pain by pinching the foot pad and observing for a reaction.</li>
		<li>Apply a veterinary eye ointment to prevent dryness during anesthesia.</li>
	</ol>
	</li>
	<li>Shave the abdominal area from the sternum down to just above the genitals using a small veterinary hair clipper. Take care to achieve a close shave without injuring the skin.</li>
	<li>Fix the rat to the surgical bed using tape on the fore- and hindlimbs, and, if using inhalation anesthesia, place the snout into a suitably-sized nose cone. Place a heating pad underneath the rat to maintain body temperature. Disinfect the abdominal area using&#160;a surgical grade disinfectant, followed by an alcohol rinse and application of an antiseptic paint.&#160;</li>
</ol>
3. Laparotomy
<ol>
	<li>Using a small curved scalpel, such as #15, make a small vertical incision in the skin layer directly along the midline. Extend the incision up to the xyphoid process. Next, incise the muscle layer by gently lifting up with a pair of Adson forceps to make a scalpel incision, and then extend the incision up to the xyphoid process with surgical scissors. Take care not to puncture the organs of the abdominal cavity or to damage the small vessels on either side of the xyphoid process.</li>
	<li>Place a retractor across the abdomen to spread the muscle and skin and expose the entire abdominal area.</li>
	<li>Moisten two 7.5 x 7.5 cm2 gauze swabs with sterile&#160;physiological saline and spread one across the top of the surgical opening and one across the bottom, as shown in Figure 2A.</li>
</ol>
4. Isolation of the Hepatic Artery
<ol>
	<li>Using two moistened cotton applicator swabs moistened with sterile physiological saline, gently lift the intestines and cecum&#160;out of the abdominal cavity, and place them on the lower gauze swab. Fold the gauze over to keep the organs moist and to hold them out of the way.</li>
	<li>Using two moistened cotton applicator swabs, gently lift up the left lateral liver lobe, which will be the most predominant lobe in view at this stage. Flip the lobe upwards, and, using a small pair of spring scissors, cut through the fibrous membrane on the underside of the lobe, allowing the lobe to be completely flipped upwards and rest on top of the previously placed gauze swab. Fold the gauze swab over the lobe to hold it in place.</li>
	<li>Continue with the next lobe (the anterior caudate), which is now the most anterior lobe in view. With the aid of a dissecting microscope and fine forceps, dissect the thin membrane surrounding the lobe in order to release it and allow it to be freely flipped upwards. Place the lobe underneath the gauze swab together with the left lateral lobe.</li>
	<li>Observe the posterior caudate lobe, which will now be visible. Repeat step 4.3 to completely dissect the posterior caudate lobe and flip it upwards and inside the gauze together with the previously placed liver lobes.</li>
	<li>Just beneath the original position of the caudate lobe, find the common hepatic artery, which will now be visible and easily recognized by its strong pulsation. Following the artery to the anatomical right, observe the thick membranous material obscuring the gastroduodenal and proper hepatic arteries from view. Using fine forceps, lift up this material and gently tear it, taking care to avoid any small vessels in this area. Use cotton applicator swabs to stop any bleeding which might occur.
	<ol>
		<li>Locate the arterial branches, which will be clearly visible with the aid of a dissecting microscope when the membranes are properly dissected.</li>
	</ol>
	</li>
	<li>Using two fine, atraumatic forceps, carefully dissect the common hepatic artery, gastroduodenal artery, and the proper hepatic artery such that they are freed from the surrounding membranes (Figure 2B).</li>
</ol>
5. Preparation of the Artery for Injection
<ol>
	<li>Pass a 7-0 Prolene suture underneath the gastroduodenal artery, and use a micro-needle-holder to tie a permanent ligature at the distal end, just above the bifurcation (see Figure 1 for proper placement). Allow approximately 2 inches of suture material remaining on the ends, and clamp them with a needle holder to pull the artery taught, as in Figure 2C.</li>
	<li>Place a small clamp on the common hepatic artery to temporarily block blood flow (Figure 2C). 
	Note: This is important to prevent massive bleeding once the needle is removed from the artery. Furthermore, the blockage of arterial blood flow will allow for slower virus administration and better transduction efficiency.
	<ol>
		<li>Important: Work quickly to limit the amount of time that blood flow is stopped - perform the injection and subsequent ligation of the artery within 5 min of clamping the common hepatic artery.</li>
	</ol>
	</li>
	<li>If possible, increase the magnification on the dissecting microscope, such that the gastroduodenal artery is in sharp focus and appears large enough to accurately insert the 30 G needle. Place the needle of the injection syringe into the lumen of the gastroduodenal artery and slowly push on the plunger. 
	Note: Proper placement of the needle will result in the injected solution flowing directly through the proper hepatic artery and towards the liver.
	<ol>
		<li>Perform the injection very slowly to maximize transduction efficiency.</li>
	</ol>
	</li>
	<li>Once the entire injection volume has been administered, slowly retract the needle from the artery. 
	Note: There should be no bleeding from the injection site. In the case that bleeding does occur, it indicates that the clamp is not properly placed and should be quickly adjusted.</li>
</ol>
6. Closing
<ol>
	<li>Pass a second 7-0 Prolene suture beneath the gastroduodenal artery, and tie a second permanent ligature above the injection site.</li>
	<li>Remove the clamp from the common hepatic artery. Confirm that blood flow through the proper hepatic artery towards the liver has been restored.
	<ol>
		<li>Cut the loose ends of the ligatures.</li>
	</ol>
	</li>
	<li>Confirm that there is no bleeding before returning the intestines and liver to the abdominal cavity. Take care that the appendix is returned in its original orientation.</li>
	<li>Suture the muscle and skin in separate layers using 4-0 vicryl suture material with a curved needle. Place continuous sutures approximately 1 mm apart and pull tightly.
	<ol>
		<li>During the suturing phase, administer an analgesic such as buprenorphine (0.05 mg/kg).</li>
	</ol>
	</li>
	<li>Remove rat from anesthesia. If isoflurane has been used, the rat should regain consciousness within several minutes. If MMF has been used as an injectable anesthesia, antagonize it with a subcutaneous injection of Atipamezole (0.75 mg/kg), Flumazenil (0.2 mg/kg), and Naloxone (9.12 mg/kg), which will reverse the effects of the anesthesia and allow the animals to regain consciousness within 20 min. Keep rats warm with an infrared warming lamp during the initial wakening phase.
	<ol>
		<li>Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
		<li>Do not return an animal that has undergone surgery to the company of other animals until fully recovered.</li>
	</ol>
	</li>
	<li>Monitor rats post-operatively for pathological signs of distress, and follow-up with analgesics as required by the local animal care and use committee. For example, buprenorphine can be administered at 0.05 mg/kg every 12 hr.</li>
</ol>

<ol>
	<li>Murray, C. J., Lopez, A. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence-based+health+policy+-+lessons+from+the+Global+Burden+of+Disease+Study.">Evidence-based health policy - lessons from the Global Burden of Disease Study.</a> Science. 274, 740-743 (1996).</li><li>Parkin, D. M., Bray, F., Ferlay, J., Pisani, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimating+the+world+cancer+burden:+Globocan+2000.">Estimating the world cancer burden: Globocan 2000.</a> Int J Cancer. 94, 153-156 (2001).</li><li>Bruix, J., Sala, M., Llovet, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemoembolization+for+hepatocellular+carcinoma.">Chemoembolization for hepatocellular carcinoma.</a> Gastroenterology. 127, 179-188 (2004).</li><li>Miraglia, R., Pietrosi, G., Maruzzelli, L., Petridis, I., Caruso, S., Marrone, G., Mamone, G., Vizzini, G., Luca, A., Gridelli, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficacy+of+transcatheter+embolization/chemoembolization+(TAE/TACE)+for+the+treatment+of+single+hepatocellular+carcinoma.">Efficacy of transcatheter embolization/chemoembolization (TAE/TACE) for the treatment of single hepatocellular carcinoma.</a> World J Gastroenterol. 13, 2952-2955 (2007).</li><li>Breedis, C., Young, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+blood+supply+of+neoplasms+in+the+liver.">The blood supply of neoplasms in the liver.</a> Am J Pathol. 30, 969-977 (1954).</li><li>Nakashima, T., Kojiro, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathologic+characteristics+of+hepatocellular+carcinoma.">Pathologic characteristics of hepatocellular carcinoma.</a> Semin Liver Dis. 6, 259-266 (1986).</li><li>Heo, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Randomized+dose-finding+clinical+trial+of+oncolytic+immunotherapeutic+vaccinia+JX-594+in+liver+cancer.">Randomized dose-finding clinical trial of oncolytic immunotherapeutic vaccinia JX-594 in liver cancer.</a> Nat Med. 19, 329-336 (2013).</li><li>Altomonte, J., Marozin, S., Schmid, R. M., Ebert, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineered+newcastle+disease+virus+as+an+improved+oncolytic+agent+against+hepatocellular+carcinoma.">Engineered newcastle disease virus as an improved oncolytic agent against hepatocellular carcinoma.</a> Mol Ther. 18, 275-284 (2010).</li><li>Shinozaki, K., Ebert, O., Kournioti, C., Tai, Y. S., Woo, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oncolysis+of+multifocal+hepatocellular+carcinoma+in+the+rat+liver+by+hepatic+artery+infusion+of+vesicular+stomatitis+virus.">Oncolysis of multifocal hepatocellular carcinoma in the rat liver by hepatic artery infusion of vesicular stomatitis virus.</a> Mol Ther. 9, 368-376 (2004).</li><li>Shinozaki, K., Ebert, O., Woo, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eradication+of+advanced+hepatocellular+carcinoma+in+rats+via+repeated+hepatic+arterial+infusions+of+recombinant+VSV.">Eradication of advanced hepatocellular carcinoma in rats via repeated hepatic arterial infusions of recombinant VSV.</a> Hepatology. 41, 196-203 (2005).</li><li>Altomonte, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synergistic+antitumor+effects+of+transarterial+viroembolization+for+multifocal+hepatocellular+carcinoma+in+rats.">Synergistic antitumor effects of transarterial viroembolization for multifocal hepatocellular carcinoma in rats.</a> Hepatology. 48, 1864-1873 (2008).</li></ol>

The authors have nothing to disclose.

Although direct intratumoral injection is undoubtedly the simplest method to result in efficient tumor transduction of a single tumor nodule, hepatic arterial infusion represents an ideal administration route to target multifocal, orthotopic HCC. This method has proven to be both safe and effective for treating HCC in immune competent rats with oncolytic viruses. Furthermore, since HCC patients are routinely treated by transarterial application of chemoembolization, the method described here is readily translatable to a clinical setting.
An obvious disadvantage of hepatic arterial delivery of tumor therapies in preclinical rodent models is the invasiveness of the procedure and the time required, both for the surgical intervention and the post-operative care of the rats. Researchers wishing to establish this technique must invest a substantial amount of time in becoming proficient at the procedure to produce reproducible results and a high survival rate. Furthermore, due to the extremely small diameter of the hepatic artery, to date there have been no reports of hepatic arterial infusions in mice. This is unfortunate, since the genetic models of HCC are primarily generated in mice, and we are therefore limited to testing hepatic arterially applied therapies in implantation or chemically induced HCC models in rats. Chemical induction of HCC is also associated with general liver toxicity, which would potentially preclude the rats from eligibility for receiving this therapeutic intervention due to safety concerns.
As an alternative to the invasive laparotomy approach to hepatic arterial virus delivery, it is theoretically feasible to feed a catheter through the femoral artery and up through the hepatic aorta and into the hepatic artery in rats, which is the standard procedure in patients. Although this approach would provide the huge benefit of avoiding surgical intervention, it is also technically challenging to establish and would require the aid of an experienced interventional radiologist.
Finally, it is noteworthy to mention that the procedure described here, although established for the purpose of efficient delivery of oncolytic viruses to orthotopic HCC, can be applied to any therapeutic agent for locoregional delivery to hepatic lesions, such as chemotherapy, immune-modulatory drugs, or small molecules. This method allows for a sophisticated pre-clinical model for testing novel therapies for clinical translation.

Hepatocellular carcinoma (HCC) is the fifth most prevalent cancer worldwide, and the third leading cause of cancer-related death, making it a significant health concern1,2. For patients who are not eligible for tumor resection, or those awaiting liver transplantation, locoregional therapy involving transarterial embolization (TAE) or transarterial chemoembolization (TACE) are applied as standard palliative care3,4. These therapies exploit the unique feature of dual blood supply in the liver whereby tumors are fed almost exclusively by hepatic arterial blood flow, while the surrounding liver receives the majority of its blood supply from the portal vein5,6.
Due to the extremely limited efficacies of established therapies for HCC, oncolytic viruses have emerged as promising alternative therapeutics. JX-594, recently renamed Pexa-Vec, is a thymidine kinase-deleted vaccinia vector, armed with granulocyte-macrophage colony-stimulating factor (GM-CSF), which has completed phase II clinical trial for HCC7. More recently, a recombinant vesicular stomatitis virus vector (VSV) expressing human interferon-beta has entered a phase I clinical trial for sorafenib-refractory HCC (NCT01628640). As oncolytic viruses move closer to obtaining approval for clinical application for HCC in patients, the need for an effective administration route to target multifocal disease is evident. While systemic delivery is largely ineffective due to inefficient tumor transduction, intratumoral applications could limit the efficacy of the therapy to the injected tumor, leaving uninjectable microscopic lesions susceptible to disease progression.
We have established a method of isolating the hepatic artery in rats to administer oncolytic virus therapy in a locoregional manner to target orthotopic HCC. We have demonstrated that this administration route results in safe and effective transduction of multifocal HCC nodules, resulting in significant survival prolongation in immune competent rats8-10. Here, we describe the method of accessing, dissecting, and injecting into the hepatic artery in rats. A scheme of the procedure is shown in Figure 1 (previously published9)

<table border="0" cellpadding="0" cellspacing="0" width="597">
	<tbody>
		<tr height="64">
			<td height="64" style="height:64px;width:144px;">Veterinary clippers</td>
			<td style="width:157px;">Aesculap</td>
			<td style="width:88px;">GT415</td>
			<td style="width:208px;">Small, cordless trimmer ideal for removing fur from surgical area</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:144px;">Stereomicroscope</td>
			<td style="width:157px;">Zeiss</td>
			<td style="width:88px;">Stemi SV6</td>
			<td style="width:208px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:144px;">30G Needles</td>
			<td style="width:157px;">Braun</td>
			<td align="right" style="width:88px;">4656300</td>
			<td style="width:208px;">30G x &#189;&#8221;</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:144px;">1ml syringes</td>
			<td style="width:157px;">Braun</td>
			<td style="width:88px;">9161406V</td>
			<td style="width:208px;">Tuberculin syringe</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:144px;">Disposable scalpel</td>
			<td style="width:157px;">Feather</td>
			<td style="width:88px;">2975#15</td>
			<td style="width:208px;">#15 blade&#160;</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Standard surgical scissors</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">14001-13&#160;</td>
			<td style="width:208px;">Sharp/blunt, for opening skin and muscle</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Adson forcep</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">1101-12</td>
			<td style="width:208px;">With teeth, for grasping skin and muscle</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:144px;">Alm retractor</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">17008-07</td>
			<td style="width:208px;">With blunt teeth, for spreading abdominal cavity open during surgery</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Gauze swabs</td>
			<td style="width:157px;">Lohmann &#38; Rauscher</td>
			<td align="right" style="width:88px;">18504</td>
			<td style="width:208px;">7.5 x 7.5 cm, should be autoclaved prior to use</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Cotton-tipped applicator swabs</td>
			<td style="width:157px;">Lohmann &#38; Rauscher</td>
			<td align="right" style="width:88px;">11970</td>
			<td style="width:208px;">Sterile</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Fine-tipped foreceps</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">11063-07</td>
			<td style="width:208px;">0.4mm, angled tip, for dissecting hepatic artery</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Vannas spring scissors</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">91500-09</td>
			<td style="width:208px;">For delicate cutting</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Micro-needle holder</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">12076-12</td>
			<td style="width:208px;">For ligating gastroduodenal artery</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:144px;">Needle holder</td>
			<td style="width:157px;">Fine Science Tools</td>
			<td style="width:88px;">12005-15</td>
			<td style="width:208px;">Tungsten carbide jaws</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:144px;">7-0 Prolene sutures</td>
			<td style="width:157px;">Ethicon</td>
			<td style="width:88px;">8648H</td>
			<td style="width:208px;">Polypropylene suture with curved needle, for ligating gastroduodenal artery</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:144px;">4-0 Vicryl sutures</td>
			<td style="width:157px;">Ethicon</td>
			<td style="width:88px;">V3040H</td>
			<td style="width:208px;">With curved needle attached</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:144px;">Infrared warming lamp</td>
			<td style="width:157px;">Beurer</td>
			<td style="width:88px;">IL11</td>
			<td style="width:208px;">For maintaining body temperature post-operatively</td>
		</tr>
	</tbody>
</table>

transarterial administration of oncolytic viruses for locoregional therapy of orthotopic hcc in rats

Hepatocellular carcinoma (HCC) is a disease with limited treatment options and poor prognosis. In recent years, oncolytic virotherapies have proven themselves to be potentially powerful tools to fight malignancy. Due to the unique dual blood supply in the liver, it is possible to apply therapies locally to orthotopic liver tumors, which are predominantly fed by arterial blood flow. We have previously demonstrated that hepatic arterial delivery of oncolytic viruses results in safe and efficient transduction efficiency of multifocal HCC lesions, resulting in significant prolongation of survival in immune competent rats. This procedure closely mimics the application of transarterial embolization in patients, which is the standard palliative care provided to many HCC patients. The ability to administer tumor therapies through the hepatic artery in rats allows for a highly sophisticated preclinical model for evaluating novel viral vectors under development. Here we describe the detailed protocol for microdissection of the hepatic artery for infusion of oncolytic virus vectors to treat orthotopic HCC.

With experience, a nearly 100% success rate (meaning that the artery was successfully dissected and injected, and the animal survived surgery) can be achieved. However, the health status of the rat prior to surgery (degree of tumor burden, underlying liver function, etc.) will obviously play a role in the outcome of the surgical intervention. Following surgery, rats can be expected to experience transient weight loss and a slight, transient elevation of liver enzymes, even if buffer alone, without virus, was injected.
We have previously demonstrated that trans-hepatic arterial administration of oncolytic vesicular stomatitis virus (VSV) or Newcastle disease virus (NDV) results in efficient tumor transduction and tumor-specific virus replication, in an orthotopic, multifocal HCC model8,9. Although the two viruses differ in their intratumoral kinetics of virus replication, at the respective time-points of maximal virus replication, numerous foci of virus propagation, as evidenced by positive enzymatic staining for the LacZ reporter gene, could be observed exclusively within HCC nodules, while the surrounding livers showed no evidence of virus replication (Figure 3 and 4, previously published data8,9). Additionally, morphometric analysis of randomly selected VSV-treated tumor nodules demonstrated that tumor transduction efficiency does not correlate with tumor size (Figure 3).
Furthermore, the ability to deliver therapeutic agents through the hepatic artery also offers the possibility to combine therapy with an embolization agent as an alternative to TACE. We have demonstrated that, by mixing oncolytic VSV at a 1:1 ratio with a degradable starch microsphere (DSM) embolization agent and infusing the suspension through the hepatic artery, we could achieve massive tumor necrosis and very significant survival prolongation in rats bearing multifocal HCC nodules11. Trans-hepatic arterial administration of the DSM resulted in a nearly complete embolization of tumor nodules, while having a minimal effect on the perfusion of normal liver tissue, as demonstrated by dynamic contrast-enhanced magnetic resonance imaging (DCE-MRI) using a gadolinium contrast agent (Figure 5, previously published data11).
<img alt="Figure 1" src="/files/ftp_upload/53757/53757fig1.jpg" /> 
Figure 1.&#160;Schematic representation of hepatic arterial infusion procedure. The hepatic vessels (common hepatic artery, proper hepatic artery, and gastroduodenal artery) are dissected with the aid of a dissecting microscope. After ligation of the gastroduodenal artery and temporal block of the common hepatic artery, 1 ml of PBS or VSV vector are administered via the gastroduodenal artery through the proper hepatic artery. After injection, the proximal site of the gastroduodenal artery is ligated to prevent bleeding, the block of the common hepatic artery is released, and the recommencement of hepatic blood flow is confirmed. This figure has been previously published9. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53757/53757fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53757/53757fig2.jpg" /> 
Figure 2. Photographs of the hepatic arterial infusion procedure. A laparotomized rat demonstrates the placement of the abdominal wall retractor and gauze swabs (A). After lifting out the hepatic lobes, the common hepatic artery (designated CHA) and gastroduodenal artery (designated GA) are identified and dissected (B). After ligating the distal end of the gastroduodenal artery, the common hepatic artery is temporarily clamped (C). The gastroduodenal artery is now ready for injection. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53757/53757fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53757/53757fig3.jpg" /> 
Figure 3. Transduction and tumor-selective replication of VSV after hepatic artery administration in rats bearing multifocal HCC. Sets of animals (n = 3/time point) were sacrificed (A) at 30 min and (B) at day 1 post-virus infusion, and frozen liver sections were stained for &#946;-gal expression. Representative sections are shown at low (left) and higher magnifications (right). Arrows indicate the borders between tumor lesions and hepatic parenchyma. (C) Relationship between tumor size and transduction efficiency. HCC-containing liver sections were analyzed by morphometric analysis to quantify total tumor and X-gal-positive areas. This figure has been previously published9. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53757/53757fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53757/53757fig4.jpg" /> 
Figure 4. NDV/F3aa(L289A) treatment results in tumor-specific virus replication in hepatocellular carcinoma (HCC)-bearing rats. Male Buffalo rats bearing multifocal orthotopic HCC nodules were treated with rNDV/F3aa(L289A) (108 TCID50) by hepatic arterial infusion, and killed at the indicated time points post-treatment (n = 3). Tumor-containing liver sections were subjected to &#946;-gal staining. Representative sections are shown at 5X (overview) and 20X (magnification of tumor and liver sections) magnification. Viral titers were quantified by TCID50 analysis of liver and tumor lysate, and are expressed as the mean &#177; SD. NDV, Newcastle disease virus; TCID50, 50% tissue culture infectious dose. Note, this Figure has been modified from its original form8. This figure has been previously published8. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53757/53757fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/53757/53757fig5.jpg" /> 
Figure 5. Dynamic contrast-enhanced magnetic resonance imaging of embolized HCC. McA-RH7777 tumor-bearing rats, either nonembolized or 30 min after hepatic arterial embolization with DSM, were imaged with DCE-MRI. (A) Axial T1-weighed precontrast (a, c) and postcontrast (b, d) images show lack of contrast in embolized tumor nodules (d). (B) Data were quantitatively analyzed by measuring gray-scale signal intensities in equal-sized regions of interest of tumor nodules and adjacent normal liver tissue. One representative data set is shown. This figure has been previously published11. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53757/53757fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Transarterial Administration of Oncolytic Viruses for Locoregional Therapy of Orthotopic HCC in Rats at JoVE.com

Transarterial Administration of Oncolytic Viruses for Locoregional Therapy of Orthotopic HCC in Rats

Oncolytic virotherapies are under development as novel therapeutics for the treatment of hepatocellular carcinoma (HCC). Here we describe a method for locoregional therapy of HCC via hepatic arterial administration of oncolytic virus.

Hepatocellular carcinoma (HCC) is a disease with limited treatment options and poor prognosis.  In recent years, oncolytic virotherapies have proven ...

transarterial-administration-oncolytic-viruses-for-locoregional

Nanyang Technological University

Research

JoVE Journal

Medicine

9.4K Views.  Klinikum rechts der Isar der TU München. Oncolytic virotherapies are under development as novel therapeutics for the treatment of hepatocellular carcinoma (HCC). Here we describe a method for locoregional therapy of HCC via hepatic arterial administration of oncolytic virus.

Video: Transarterial Administration of Oncolytic Viruses for Locoregional Therapy of Orthotopic HCC in Rats

Ex Vivo Infection of Live Tissue with Oncolytic Viruses

Oncolytic Viruses (OVs) are novel therapeutics that selectively replicate in and kill tumor cells1. Several clinical trials evaluating the effectiveness of a variety of oncolytic platforms including HSV, Reovirus, and Vaccinia OVs as treatment for cancer are currently underway2-5. One key characteristic of oncolytic viruses is that they can be genetically modified to express reporter transgenes which makes it possible to visualize the infection of tissues by microscopy or bio-luminescence imaging6,7. This offers a unique advantage since it is possible to infect tissues from patients ex vivo prior to therapy in order to ascertain the likelihood of successful oncolytic virotherapy8. To this end, it is critical to appropriately sample tissue to compensate for tissue heterogeneity and assess tissue viability, particularly prior to infection9. It is also important to follow viral replication using reporter transgenes if expressed by the oncolytic platform as well as by direct titration of tissues following homogenization in order to discriminate between abortive and productive infection. The object of this protocol is to address these issues and herein describes 1. The sampling and preparation of tumor tissue for cell culture 2. The assessment of tissue viability using the metabolic dye alamar blue 3. Ex vivo infection of cultured tissues with vaccinia virus expressing either GFP or firefly luciferase 4. Detection of transgene expression by fluorescence microscopy or using an In Vivo Imaging System (IVIS) 5. Quantification of virus by plaque assay. This comprehensive method presents several advantages including ease of tissue processing, compensation for tissue heterogeneity, control of tissue viability, and discrimination between abortive infection and bone fide viral replication.

Ex Vivo Infection of Live Tissue with Oncolytic Viruses

Oncolytic viruses are promising for cancer therapeutics. The ability to ascertain the infectability of live tissue specimens obtained from patients prior to treatment is a unique advantage of this therapeutic approach. This protocol describes how to process tissues for ex vivo infection with oncolytic virus and subsequent viral quantification.

Oncolytic Viruses (OVs) are novel therapeutics that selectively replicate in and kill tumor cells1. Several clinical trials evaluating the effectiveness ...

Multi-photon Imaging of Tumor Cell Invasion in an Orthotopic Mouse Model of Oral Squamous Cell Carcinoma

Loco-regional invasion of head and neck cancer is linked to metastatic risk and presents a difficult challenge in designing and implementing patient management strategies. Orthotopic mouse models of oral cancer have been developed to facilitate the study of factors that impact invasion and serve as model system for evaluating anti-tumor therapeutics. In these systems, visualization of disseminated tumor cells within oral cavity tissues has typically been conducted by either conventional histology or with in vivo bioluminescent methods. A primary drawback of these techniques is the inherent inability to accurately visualize and quantify early tumor cell invasion arising from the primary site in three dimensions. Here we describe a protocol that combines an established model for squamous cell carcinoma of the tongue (SCOT) with two-photon imaging to allow multi-vectorial visualization of lingual tumor spread. The OSC-19 head and neck tumor cell line was stably engineered to express the F-actin binding peptide LifeAct fused to the mCherry fluorescent protein (LifeAct-mCherry). Fox1nu/nu mice injected with these cells reliably form tumors that allow the tongue to be visualized by ex-vivo application of two-photon microscopy. This technique allows for the orthotopic visualization of the tumor mass and locally invading cells in excised tongues without disruption of the regional tumor microenvironment. In addition, this system allows for the quantification of tumor cell invasion by calculating distances that invaded cells move from the primary tumor site. Overall this procedure provides an enhanced model system for analyzing factors that contribute to SCOT invasion and therapeutic treatments tailored to prevent local invasion and distant metastatic spread. This method also has the potential to be ultimately combined with other imaging modalities in an in vivo setting.

A comprehensive overview of the techniques involved in generating a mouse model of oral cancer and quantitative monitoring of tumor invasion within the tongue through multi-photon microscopy of labeled cells is presented. This system can serve as a useful platform for the molecular assessment and drug efficacy of anti-invasive compounds.

Loco-regional invasion of head and neck cancer is linked to metastatic risk and presents a difficult challenge in designing and implementing patient ...

Production and Detection of Reactive Oxygen Species (ROS) in Cancers

Reactive oxygen species include a number of molecules that damage DNA and RNA and oxidize proteins and lipids (lipid peroxydation). These reactive molecules contain an oxygen and include H2O2 (hydrogen peroxide), NO (nitric oxide), O2- (oxide anion), peroxynitrite (ONOO-), hydrochlorous acid (HOCl), and hydroxyl radical (OH-).
Oxidative species are produced not only under pathological situations (cancers, ischemic/reperfusion, neurologic and cardiovascular pathologies, infectious diseases, inflammatory diseases 1, autoimmune diseases 2, etc&hellip;) but also during physiological (non-pathological) situations such as cellular metabolism 3, 4. Indeed, ROS play important roles in many cellular signaling pathways (proliferation, cell activation 5, 6, migration 7 etc..). ROS can be detrimental (it is then referred to as "oxidative and nitrosative stress") when produced in high amounts in the intracellular compartments and cells generally respond to ROS by upregulating antioxidants such as superoxide dismutase (SOD) and catalase (CAT), glutathione peroxidase (GPx) and glutathione (GSH) that protects them by converting dangerous free radicals to harmless molecules (i.e. water). Vitamins C and E have also been described as ROS scavengers (antioxidants).
Free radicals are beneficial in low amounts 3. Macrophage and neutrophils-mediated immune responses involve the production and release of NO, which inhibits viruses, pathogens and tumor proliferation 8. NO also reacts with other ROS and thus, also has a role as a detoxifier (ROS scavenger). Finally NO acts on vessels to regulate blood flow which is important for the adaptation of muscle to prolonged exercise 9, 10. Several publications have also demonstrated that ROS are involved in insulin sensitivity 11, 12.
Numerous methods to evaluate ROS production are available. In this article we propose several simple, fast, and affordable assays; these assays have been validated by many publications and are routinely used to detect ROS or its effects in mammalian cells. While some of these assays detect multiple ROS, others detect only a single ROS.

Production and Detection of Reactive Oxygen Species ROS in Cancers

Here we propose simple methods to test and evaluate the presence of reactive oxygen species in cells.

Reactive oxygen species include a number of molecules that damage DNA and RNA and oxidize proteins and lipids (lipid peroxydation). These reactive ...

Orthotopic Small Bowel Transplantation in Rats

Small bowel transplantation has become an accepted clinical option for patients with short gut syndrome and failure of parenteral nutrition (irreversible intestinal failure). In specialized centers improved operative and managing strategies have led to excellent short- and intermediate term patient and graft survival while providing high quality of life 1,3. Unlike in the more common transplantation of other solid organs (i.e. heart, liver) many underlying mechanisms of graft function and immunologic alterations induced by intestinal transplantation are not entirely known6,7. Episodes of acute rejection, sepsis and chronic graft failure are the main obstacles still contributing to less favorable long term outcome and hindering a more widespread employment of the procedure despite a growing number of patients on home parenteral nutrition who would potentially benefit from such a transplant. The small intestine contains a large number of passenger leucocytes commonly referred to as part of the gut associated lymphoid system (GALT) this being part of the reason for the high immunogenity of the intestinal graft. The presence and close proximity of many commensals and pathogens in the gut explains the severity of sepsis episodes once graft mucosal integrity is compromised (for example by rejection). To advance the field of intestinal- and multiorgan transplantation more data generated from reliable and feasible animal models is needed. The model provided herein combines both reliability and feasibility once established in a standardized manner and can provide valuable insight in the underlying complex molecular, cellular and functional mechanisms that are triggered by intestinal transplantation. We have successfully used and refined the described procedure over more than 5 years in our laboratory 8-11. The JoVE video-based format is especially useful to demonstrate the complex procedure and avoid initial pitfalls for groups planning to establish an orthotopic rodent model investigating intestinal transplantation.

Small bowel transplantation has become an accepted treatment option for patients with irreversible intestinal failure. Our experimental model of orthotopic small bowel transplantation in rats serves as a reliable tool to address underlying immunologic and inflammatory processes that complicate intestinal transplantation.

Small  bowel transplantation has become an accepted clinical option for patients with  short gut syndrome and failure of parenteral nutrition ...

Orthotopic Liver Transplantation in Rats

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat liver transplantation in 1979 by Kamada et al.3, this model has remained the gold standard despite various proposed alternative techniques4. Nevertheless, its broader use is limited by its steep learning curve5.
In this video paper, we show a simple and easy-to-establish revision of Kamada's two-cuff technique. The suprahepatic vena cava anastomosis is performed manually with a running suture, and the vena porta and infrahepatic vena cava anastomoses are performed utilizing a quick-linker cuff system6. Manufacturing the quick-linker kit is shown in a separate video paper.

We present an easy-to-establish revision of the classical two-cuff technique for orthotopic liver transplantation in rat.

Clinical progress in the field of liver transplantation has been largely supported by animal models1,2. Since the publication of the first orthotopic rat ...

A Novel Surgical Approach for Intratracheal Administration of Bioactive Agents in a Fetal Mouse Model

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and acquired diseases. Apart from congenital or inherited abnormalities with the requirement for long-term expression of the delivered gene, several non-inherited perinatal conditions, where short-term gene expression or pharmacological intervention is sufficient to achieve therapeutic effects, are considered as potential future indications for this kind of approach. Candidate diseases for the application of short-term prenatal therapy could be the transient neonatal deficiency of surfactant protein B causing neonatal respiratory distress syndrome1,2 or hyperoxic injuries of the neonatal lung3. Candidate diseases for permanent therapeutic correction are Cystic Fibrosis (CF)4, genetic variants of surfactant deficiencies5 and &alpha;1-antitrypsin deficiency6.
Generally, an important advantage of prenatal gene therapy is the ability to start therapeutic intervention early in development, at or even prior to clinical manifestations in the patient, thus preventing irreparable damage to the individual. In addition, fetal organs have an increased cell proliferation rate as compared to adult organs, which could allow a more efficient gene or stem cell transfer into the fetus. Furthermore, in utero gene delivery is performed when the individual's immune system is not completely mature. Therefore, transplantation of heterologous cells or supplementation of a non-functional or absent protein with a correct version should not cause immune sensitization to the cell, vector or transgene product, which has recently been proven to be the case with both cellular and genetic therapies7.
In the present study, we investigated the potential to directly target the fetal trachea in a mouse model. This procedure is in use in larger animal models such as rabbits and sheep8, and even in a clinical setting9, but has to date not been performed before in a mouse model. When studying the potential of fetal gene therapy for genetic diseases such as CF, the mouse model is very useful as a first proof-of-concept because of the wide availability of different transgenic mouse strains, the well documented embryogenesis and fetal development, less stringent ethical regulations, short gestation and the large litter size.
Different access routes have been described to target the fetal rodent lung, including intra-amniotic injection10-12, (ultrasound-guided) intrapulmonary injection13,14 and intravenous administration into the yolk sac vessels15,16 or umbilical vein17. Our novel surgical procedure enables researchers to inject the agent of choice directly into the fetal mouse trachea which allows for a more efficient delivery to the airways than existing techniques18.

We developed a novel surgical approach for intratracheal administration of bioactive agents into the mouse fetus. The delivery route is more efficient in targeting the fetal mouse lungs than the commonly used intra-amniotic injection. This procedure has to date not been described in a mouse model.

Prenatal pulmonary delivery of cells, genes or pharmacologic agents could provide the basis for new therapeutic strategies for a variety of genetic and ...

Cytotoxic Efficacy of Photodynamic Therapy in Osteosarcoma Cells In Vitro

In recent years, there has been the difficulty in finding more effective therapies against cancer with less systemic side effects. Therefore Photodynamic Therapy is a novel approach for a more tumor selective treatment.
Photodynamic Therapy (PDT) that makes use of a nontoxic photosensitizer (PS), which, upon activation with light of a specific wavelength in the presence of oxygen, generates oxygen radicals that elicit a cytotoxic response1. Despite its approval almost twenty years ago by the FDA, PDT is nowadays only used to treat a limited number of cancer types (skin, bladder) and nononcological diseases (psoriasis, actinic keratosis)2.
The major advantage of the use of PDT is the ability to perform a local treatment, which prevents systemic side effects. Moreover, it allows the treatment of tumors at delicate sites (e.g. around nerves or blood vessels). Here, an intraoperative application of PDT is considered in osteosarcoma (OS), a tumor of the bone, to target primary tumor satellites left behind in tumor surrounding tissue after surgical tumor resection. The treatment aims at decreasing the number of recurrences and at reducing the risk for (postoperative) metastasis.
In the present study, we present in vitro PDT procedures to establish the optimal PDT settings for effective treatment of widely used OS cell lines that are used to reproduce the human disease in well established intratibial OS mouse models. The uptake of the PS mTHPC was examined with a spectrophotometer and phototoxicity was provoked with laser light excitation of mTHPC at 652 nm to induce cell death assessed with a WST-1 assay and by the counting of surviving cells. The established techniques enable us to define the optimal PDT settings for future studies in animal models. They are an easy and quick tool for the evaluation of the efficacy of PDT in vitro before an application in vivo.

Cytotoxic Efficacy of Photodynamic Therapy in Osteosarcoma Cells In Vitro

The protocol reported here allows the evaluation of the efficacy of photodynamic therapy (PDT) in cell lines and the optimization of the PDT settings before applying the therapy in animal models.

In recent years, there has been the difficulty in finding more effective therapies against cancer with less systemic side effects. Therefore Photodynamic ...

Handling of the Cotton Rat in Studies for the Pre-clinical Evaluation of Oncolytic Viruses

Oncolytic viruses are a novel anticancer therapy with the ability to target tumor cells, while leaving healthy cells intact. For this strategy to be successful, recent studies have shown that involvement of the host immune system is essential. Therefore, oncolytic virotherapy should be evaluated within the context of an immunocompetent model. Furthermore, the study of antitumor therapies in tolerized animal models may better recapitulate results seen in clinical trials. Cotton rats, commonly used to study respiratory viruses, are an attractive model to study oncolytic virotherapy as syngeneic models of mammary carcinoma and osteosarcoma are well established. However, there is a lack of published information on the proper handling procedure for these highly excitable rodents. The handling and capture approach outlined minimizes animal stress to facilitate experimentation. This technique hinges upon the ability of the researcher to keep calm during handling and perform procedures in a timely fashion. Finally, we describe how to prepare cotton rat mammary tumor cells for consistent subcutaneous tumor formation, and how to perform intratumoral and intraperitoneal injections. These methods can be applied to a wide range of studies furthering the development of the cotton rat as a relevant pre-clinical model to study antitumor therapy.

Cotton rats are extremely excitable and have a strong flight-or-fight response. A handling method optimized to reduce the stress of the animals is described which will make cotton rats more accessible as a preclinical model.

Oncolytic viruses are a novel anticancer therapy with the ability to target tumor cells, while leaving healthy cells intact.  For this strategy to be ...

Design, Fabrication, and Administration of the Hand Active Sensation Test (HASTe)

The concept of personalizing neurologic rehabilitation, based on individual impairments, has experienced a recent surge. In parallel, the number of outcome measures of upper extremity motor performance has grown. However, clinicians and researchers lack practical, quantitative measures of the hand&#8217;s natural role as a receptor of the environment. The Hand Active Sensation Test (HASTe), developed by Williams and colleagues in 2006, is a valid and reliable measure of haptic performance. Though not available commercially, the HASTe can be fabricated from inexpensive materials, and it has been used successfully to identify impairments in haptic touch in individuals with stroke. (Williams, 2006). This paper presents the methods of design and fabrication of the HASTe testing kit, as well as a visual screen to be used during administration, and instructions for the tests administration and scoring.

Design, Fabrication, and Administration of the Hand Active Sensation Test HASTe

The Hand Active Sensation Test (HASTe) is a valid and reliable measure of haptic performance, which has been used successfully to identify impaired haptic touch in individuals with stroke. The purpose of this paper is to describe the design, fabrication and administration of the HASTe.

The concept of personalizing neurologic rehabilitation, based on individual impairments, has experienced a recent surge. In parallel, the number of ...

A Method for Quantifying Upper Limb Performance in Daily Life Using Accelerometers

A key reason for referral to rehabilitation services after stroke and other neurological conditions is to improve one's ability to function in daily life. It has become important to measure a person's activities in daily life, and not just measure their capacity for activity in the structured environment of a clinic or laboratory. A wearable sensor that is now enabling measurement of daily movement is the accelerometer. Accelerometers are commercially-available devices resembling large wrist watches that can be worn throughout the day. Data from accelerometers can quantify how the limbs are engaged to perform activities in peoples' homes and communities. This report describes a methodology to collect accelerometry data and turn it into clinically-relevant information. First, data are collected by having the participant wear two accelerometers (one on each wrist) for 24 h or longer. The accelerometry data are then downloaded and processed to produce four different variables that describe key aspects of upper limb activity in daily life: hours of use, use ratio, magnitude ratio, and the bilateral magnitude. Density plots can be constructed that visually represent the data from the 24 h wearing period. The variables and their resultant density plots are highly consistent in neurologically-intact, community-dwelling adults. This striking consistency makes them a useful tool for determining if upper limb daily performance is different from normal. This methodology is appropriate for research studies investigating upper limb dysfunction and interventions designed to improve upper limb performance in daily life in people with stroke and other patient populations. Because of its relative simplicity, it may not be long before it is also incorporated in clinical neurorehabilitation practice.

This protocol describes a method to quantify upper limb performance in daily life using wrist-worn accelerometers.

A key reason for referral to rehabilitation services after stroke and other neurological conditions is to improve one's ability to function in daily life. ...