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EoE Sub Articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Creation of in vitro VX2 cell line
<ol>
	<li>Harvest the VX2 tumor from rabbit quadricep muscle and growth in mouse flank.
	<ol>
		<li>Thaw frozen VX2 tumor blocks (1 cm x 1 cm), mince on a culture plate using a scalpel blade and strain through a 70 &#181;m filter by adding small amounts of Hanks Balanced Salt Solution (HBSS) sequentially to ensure all cells are strained for a final volume of 1 mL.</li>
		<li>Count cells and dilute with 0.9% phosphate buffered saline solution to a concentration of 1 x 107/mL and place on ice in a sterile tube. 
		NOTE: Tumor blocks were obtained from a lab collaborator from previous propagation of VX2 tumors in rabbit quadriceps muscles.</li>
		<li>Anesthetize a female white New Zealand rabbit (weight 2.5-3.5 kg) with 2-5% inhaled isoflurane, remove the fur overlying the injection site and cleanse the injection site with povidone-iodine solution. Inject 500 &#181;L of the prepared VX2 cell suspension (5 x 106&#160;cells/mL) into the quadriceps muscle of the rabbit. Monitor the rabbit clinically for tumor growth starting on day 10.</li>
		<li>Euthanize the rabbits after tumors reach 1-1.5 cm in diameter (measured externally with calipers) using&#160;a veterinarian approved method. Confirm euthanasia by checking vital signs including heart rate and respiratory rate. Excise the tumor using sterile instruments after cleaning the area with betadine solution.</li>
		<li>In a biological safety cabinet using sterile instruments, mince the tumor into small pieces (0.2 cm) on a 10 cm tissue culture dish using a scalpel blade. Pass the tumor fragments through a 70 &#181;m cell strainer using 1 mL of HBSS as described in step 1.1.1. Count cells and dilute using 0.9% saline solution to obtain 7.5 x 105&#160;cells in 200 &#181;L.</li>
		<li>Anesthetize male NOD SCID gamma mice (weight 25 g) using 4% isoflurane at 1 L/min and maintain anesthesia using 2% isoflurane. Once mice are anesthetized, inject 200 &#181;L of the solution from step 1.1.5 into the subcutaneous tissue of the mouse flank using a 27 gauge needle.</li>
	</ol>
	</li>
	<li>Harvesting and passaging of VX2 cells in vitro
	<ol>
		<li>Monitor xenograft growth clinically twice weekly until tumors reach 1 cm in diameter using calipers.</li>
		<li>Euthanize mice by placing in a CO2&#160;chamber or performing cervical dislocation after anesthetizing with 4% isoflurane gas. Excise tumors under sterile conditions in a biological safety cabinet.</li>
		<li>Mince tumors with a scalpel into 1-2 mm pieces in a 6 cm tissue culture dish containing 3 mL of Dulbecco&#39;s Modified Eagle Medium (DMEM)/Ham&#39;ss F12+ 10% fetal bovine serum (FBS) media.</li>
		<li>Leave tumor fragments undisturbed in a 37 &#176;C incubator with 5% CO2&#160;for two days until cells begin to grow out from the tumor pieces. Subsequently, check cultured plates daily by light microscopy for cell confluency. Upon reaching 70% confluency, trypsinize cells by adding 1 mL of 0.05% trypsin to the flask and placing back in the 37 &#176;C incubator for 5 min.</li>
		<li>Neutralize trypsin with 6 mL of DMEM/Ham&#39;ss F12 + 10% FBS medium, collect cells and centrifuge at 4 &#176;C for 5 min at 300 x&#160;g. Remove supernatant and re-suspend pellet in 3.5 mL of new DMEM/Ham&#39;ss F12 + 10% FBS media. Seed fresh 6 cm collagen I coated plates with 1.6 x 106&#160;cells per plate to achieve approximately 50% cell seeding density. 
		&#8203;NOTE: 50% cell density refers to the cells taking up 50% of the surface area of the plate when attached.</li>
		<li>Early passaging:&#160;Repeat the above process (trypsinizing, seeding) until cells have been passaged 5 times and then assess cell line purity with PCR using a quantitative PCR assay to distinguish rabbit genomic DNA from mouse genomic DNA (see step 1.3.1-1.3.4). 
		NOTE: After 8 passages, cells can be propagated on non-collagen coated regular adherent tissue culture plates.</li>
		<li>Passage, trypsinize and freeze aliquots of cells in DMEM/HAM F12+30% FBS+10%DMSO at a concentration of 2 x 106&#160;per vial. Store cells in liquid nitrogen until required for experiments.</li>
	</ol>
	</li>
	<li>Confirmation of VX2 origin of surviving cells:
	<ol>
		<li>Create a standard cure from purified genomic mouse and rabbit DNA (Step 1.3.2 and Step 1.3.3). Use primers which target species-specific sequences within the second open reading frame of the LINE-1 retrotransposon element. 
		NOTE: Primer sequences for CRPV E6 are: 5&#39;s- GATCCTGGACCCAACCAGTGand 5&#39;s-CCTGCCGGTCCCTGATTTAT. The LINE-1 retrotransposon elements were used. Primer sequences for rabbit LINE-1 are: 5&#39;s-TCAGGAAACCCCAGAAAGTATGC and 5&#39;s-TTTGATTTCTTGAATGACCCAGTGT Primer sequences for mouse LINE-1 are: 5&#39;s-AATGGAAAGCCAACATTCACGTG and 5&#39;s-CCTTCCTTGACCAAGGTATCATTG</li>
		<li>To create a standard curve for CRPV E6, purify VX2 tumor cell lysate (Step 1.1.1) using a commercial kit following manufacturer&#39;ss protocol. Quantify this DNA using a commercial assay. Perform 6 serial dilutions from 225 pg/&#181;L to 0.925 pg/&#181;L in low EDTA-TE buffer. To create a standard curve for mouse genomic DNA, dilute 100 &#181;g of commercial mouse genomic DNA in low EDTA-TE buffer in 5 dilutions from 125 pg/&#181;L down to 0.0125 pg/&#181;L.</li>
		<li>Place 4 &#181;L from each diluted solution from the samples from step 1.3.2 in wells and run samples in a PCR thermocycler. Use the Cq values from each run along with the DNA amounts per well to calculate a standard curve using the thermocycler software auto-threshold settings.</li>
		<li>Use standard qPCR procedures to perform PCR with 40 cycles of 2-step cycling (98 &#176;C and 60 &#176;C) on a thermocycler. Establish threshold values, and set delta Ct values at &#62;35 to ensure there is minimal mouse contamination in the rabbit VX2 cell line. The total volume of each reaction was 10 &#181;L, consisting of 5 &#181;L of 2x Mastermix, 1 &#181;L of forward + reverse primers with final primer concentration in reaction at 250 nM, and 4 &#181;L of DNA sample. 
		NOTE: Threshold values are established, and delta Ct values are set at &#62;35 to ensure there is minimal mouse contamination in the rabbit VX2 cell line. The recommended level of VX2 DNA for reliable detection by qPCR is 1-10 pg.</li>
	</ol>
	</li>
</ol>
2. VX2 cell culture and creation of cell suspension
<ol>
	<li>Thaw vials containing VX2 cells (frozen in step 1.2.7) in a water bath at 37 &#176;C for 1 minute and transfer cells to a conical centrifuge tube with 10 mL of culture medium (1:1 DMEM/F-12 + 10% FBS and 1% Penicillin-streptomycin). Centrifuge for 8 min at 107 x&#160;g, discard the supernatant and re-suspend the cell pellet in 9 mL of media.</li>
	<li>Transfer re-suspended cells to a large culture flask. Incubate cells without shaking at 37 &#176;C. Check cells daily for confluence using light microscopy and change culture media every three days.</li>
	<li>Creation of cultured VX2 cell suspension for injection
	<ol>
		<li>Trypsinize cells by adding 3 mL of 0.25% trypsin to flask once cells have reached 80% confluence. Place flask in incubator (37 &#176;C) for 5 min. Transfer solution to a conical centrifuge tube and centrifuge for 8 min at 107 x&#160;g&#160;and then remove supernatant.</li>
		<li>Wash the cell pellets 3 times with 9 mL of phosphate buffered saline, centrifuge and remove supernatant as above. Count the cells and dilute with 0.9% phosphate buffered saline solution to a concentration of 4 x 107&#160;cells/mL and place in a sterile conical centrifuge tube on ice.</li>
	</ol>
	</li>
	<li>Creation of in vivo propagated VX2 cell suspension for injection
	<ol>
		<li>Thaw frozen VX2 tumor blocks (1 cm x 1 cm), mince on a culture plate using a scalpel blade and strain through a 70 &#181;m filter as in step 1.1.1. Add small amounts of HBSS sequentially to ensure all cells are strained for a final volume of 1 mL. Count cells and dilute with 0.9% phosphate buffered saline to a concentration of 1 x107/mL and place on ice in a sterile tube. 
		NOTE: Tumor blocks were obtained from a lab collaborator from previous propagation of VX2 tumors in rabbit quadriceps muscles.</li>
	</ol>
	</li>
</ol>
3. Surgical Model Establishment
<ol>
	<li>Establishment of surgical anesthesia and pre-operative preparation
	<ol>
		<li>Pre-medicate female white New Zealand rabbits (2.5-3.5 kg) 1 h&#160;prior to the planned surgical procedure using injections of acepromazine (1 mg/kg IM) and meloxicam (0.2 mg/kg SQ).&#160;Anesthetize a female white New Zealand rabbit (weight 2.5-3.5 kg) with 2-5% inhaled isoflurane, remove the fur overlying the injection site and cleanse the injection site with povidone-iodine solution.</li>
		<li>Intubate anesthetized rabbits using a laryngeal mask airway (LMA) and secure in place with tape, maintaining deep anesthesia by titrating the inhaled isoflurane dose between 2-5%. Monitor surgical anesthesia regularly throughout the procedure by checking vital signs (respiratory rate, capillary refill, oxygen saturation if available) and monitoring for signs suggestive of pain (movement, withdrawal from stimuli, noises) or light anesthesia (movement, chewing on LMA tube). 
		NOTE: This should be done by someone with experience monitoring surgical anesthesia.</li>
		<li>Place a 22-gauge ear-vein catheter in the marginal dorsal vein. Administer cefazolin (20 mg/kg) intravenously 10 min prior to surgical skin incision.</li>
		<li>Clip hair over the pelvis and abdomen prior to placing the rabbit on the operating table, then place rabbits in a dorsal position on the operating table. Clean the surgical field using a 3-step surgical skin prep (betadine soap, chlorhexidine solution, betadine solution). Drape the surgical field with laparotomy drapes after land-marking the top of the pubic bone, leaving a 5 cm x 5 cm area of the lower abdomen exposed.</li>
	</ol>
	</li>
	<li>Creation of the laparotomy incision and identification of the uterine horns
	<ol>
		<li>Put on a surgical cap and face mask. Scrub hands using chlorhexidine or betadine surgical scrub solution. Using sterile technique, put on a sterile gown and sterile surgical gloves.</li>
		<li>Using a # 11-blade scalpel, make a 2.5 cm long incision 1 cm cranial to the symphysis pubis through skin and subcutaneous tissue of the rabbit abdomen. Incise the rectus fascia and dissect the rectus muscles laterally to expose the underlying peritoneum. Enter the peritoneum sharply after ensuring the under surface is clear of bowel or other abdominal organs.</li>
		<li>Locate the uterine horns identifying the urinary bladder and sweeping a gloved finger superiorly, posteriorly and laterally over the apex of the bladder. 
		NOTE: If necessary, the full bladder can be emptied using digital pressure. Once located, bring the uterine horns through the abdominal incision to rest on the abdominal wall.</li>
		<li>Using a 3-0 braided absorbable suture, perform a single suture ligation of each uterine horn approximately 1.5-2.0 cm distal to the cervices. Place the suture just medial to the uterine arteries, which run along the lateral aspect of each horn. Tie the sutures snugly to occlude the distal uterine horns (Figure 1).</li>
	</ol>
	</li>
	<li>Myometrial VX2 inoculation
	<ol>
		<li>Using a 27-gauge needle, inject 0.5 mL of the previously prepared VX2 cell suspension from either step 2.3.2 (for cultured VX2 model) or 2.4 (for in vivo propagated VX2 model) into the myometrium of each uterine horn proximal to the suture site (between the suture site and the cervix). Inject over 1 minute and ensure that cells are not being injected into the underlying uterine cavity.&#160;Anesthetize animals using inhaled isoflurane (2-5%) and an anesthetic machine with a Bain circuit.</li>
		<li>Apply pressure to the myometrial injection site for 30 s after injection to minimize leakage of cells. Inspect the injection and suture sites for hemostasis and place the uterine horns back into the abdomen.</li>
	</ol>
	</li>
	<li>Closing the surgical incision
	<ol>
		<li>Deep abdominal wall closure: Identify the apex of the peritoneal incision and grasp the peritoneum, rectus muscle and fascia with tissue forceps.
		<ol>
			<li>To do this, anchor the suture at one apex of the incision by suturing superficial to deep on one side of the incision and deep to superficial on the other. Tie a knot. Using the attached suture, make running stitches perpendicular to the incision through the layers of the abdominal wall working step-wise along the incision from side to side. Tie the suture and cut.</li>
		</ol>
		</li>
		<li>Superficial abdominal wall closure: Identify the apex of the skin incision and suture the abdominal skin using buried running subcuticular 3-0 absorbable poly-filament suture. Apply surgical glue to the closed incision to oppose the skin edges.
		<ol>
			<li>To do this, anchor the suture at one apex of the incision, by suturing deep to superficial on one side of the incision and superficial to deep on the other. Tie a knot. Using the attached sutures, make running stitches in the dermal layer parallel to the incision working step-wise along the incision from side to side. Tie the suture and cut.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Post-operative care
	<ol>
		<li>Awakening from anesthesia: Turn off the isoflurane once the incision is closed and let the rabbit breath oxygen by mask while monitoring for signs of awakening (e.g., spontaneous movements, eye opening and chewing motions on the laryngeal mask airway). Extubate the rabbit once these signs are noted and provide&#160;oxygen by mask and a warm blanket until the rabbits are alert and able to sit independently.</li>
		<li>Post-operative monitoring: Monitor rabbits twice a day for 4 days and daily for an additional 10 days post-operatively. To monitor, assess their general condition, their food and water intake, urine and fecal output, pain assessment, weight, vital signs (heart rate, respiration rate) and their surgical site looking for swelling, erythema, discharge or dehiscence.</li>
		<li>Post-operative pain control: Administer Meloxicam daily (0.2 mg/kg SQ) for 48 h&#160;post-operatively and then as needed based on pain assessment.&#160;Administer buprenorphine immediately post-operatively and every 12 h&#160;(0.01-0.05 mg/kg SQ) for 24 h and then as needed based on pain assessment</li>
		<li>Antibiotics (enrofloxacin 5mg/kg IM) may be administered daily for seven days post-operatively to prevent wound infection as recommended by your veterinarian.&#160;Administer subcutaneous fluids (15 mL of SQ) twice a day as needed for dehydration and soft foods or other dietary supplements are provided daily as needed for weight loss.</li>
	</ol>
	</li>
</ol>

<table><tbody><tr><td>11-blade scalpel, Sterile, Disposible</td><td>Aspen Surgical (VWR)</td><td>80094-086</td><td></td></tr><tr><td>22-gague ear vein catheter</td><td>CDMV</td><td>14332</td><td></td></tr><tr><td>3-0 absorbable poly-filament suture (Polysorb)</td><td>Covidien</td><td>356718</td><td></td></tr><tr><td>3-0 braided absorbable suture (Polysorb)</td><td>Covidien</td><td>356718</td><td></td></tr><tr><td>70uM cell strainer, Individually wrapped, Nylon</td><td>Falcon</td><td>352350</td><td></td></tr><tr><td>Acepromazine (Atravet)</td><td>CDMV</td><td>1047</td><td></td></tr><tr><td>Betadine soap (Poviodone iodine 7.5%)</td><td>CDMV</td><td>4363</td><td></td></tr><tr><td>Betadine solution (Poviodone iodine 10%)</td><td>UHN Stores</td><td>457955</td><td></td></tr><tr><td>Buprenorphine</td><td>McGill University</td><td></td><td></td></tr><tr><td>Cefazolin</td><td>UHN in-patient pharmacy</td><td>No Cat # Needed</td><td></td></tr><tr><td>Chlorhexidine solution</td><td>CDMV</td><td>119872</td><td></td></tr><tr><td>Corning BioCoatCellware, Collagen Type I, 100mm dishes</td><td>Corning</td><td>354450</td><td>Brand not important</td></tr><tr><td>Corning BioCoatCellware, Collagen Type I, 24-well plates</td><td>Corning</td><td>354408</td><td>Brand not important</td></tr><tr><td>Corning BioCoatCellware, Collagen Type I, 6-well plates</td><td>Corning</td><td>354400</td><td>Brand not important</td></tr><tr><td>Corning Matrigel Basement Membrane Matrix, *LDEV-free, 10 mL</td><td>Corning</td><td>354234</td><td></td></tr><tr><td>DMEM/HAM F12 1:1</td><td>Life Technologies</td><td>11320</td><td>Brand not important</td></tr><tr><td>DMSO</td><td>Caledon Lab Chem</td><td>10/1/4100</td><td></td></tr><tr><td>Enrofloxacin (Baytril injectable)</td><td>CDMV</td><td>11242</td><td></td></tr><tr><td>Falcon Tube</td><td>Corning Centri-Star</td><td>430828</td><td></td></tr><tr><td>Fetal Bovine Serum, Qualified, Canadian Origin, 500ml</td><td>Life Technologies</td><td>12483020</td><td>Brand/source not important</td></tr><tr><td>Isoflurane</td><td>UHN in-patient pharmacy</td><td>No Cat # Needed</td><td></td></tr><tr><td>Isohexol contrast</td><td>GE Healthcare</td><td>407141210</td><td></td></tr><tr><td>Meloxicam (Metacam 0.5%)</td><td>CDMV</td><td>104674</td><td></td></tr><tr><td>Normal Saline</td><td>House Brand (UofT Medstore)</td><td>1011</td><td></td></tr><tr><td>PBS</td><td>Multicell or Sigma</td><td>331-010-CL or D8537-500mL</td><td></td></tr><tr><td>Penicillin/Streptomycin (100mL; 10000U Penicillin, 10000ug Streptomycin)</td><td>Corning-Cellgro</td><td>CA45000-652</td><td></td></tr><tr><td>Sterile Hanks Balanced Salt Solution (-Ca++, -Mg++, -Phenol Red)</td><td>T.C.M.F (Dr Bristow)</td><td>28-Jan-11</td><td></td></tr><tr><td>Surgical Glue (Tissue Adhesive)</td><td>3M Vetbond</td><td>14695B</td><td></td></tr><tr><td>Trypsin (0.25%), Proteomics Grade</td><td>Sigma</td><td>T-6567-5X20UG</td><td></td></tr><tr><td>Trypsin-EDTA, 0.05%, 100ml</td><td>Wisent Inc</td><td>325-542-EL</td><td>Brand not important</td></tr></tbody></table>

modeling metastatic endometrial cancer  model: a technique to generate endometrial cancer with lymph node metastasis in rabbit models

Source: Lauren Philp et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/t/59340">An Orthotopic Endometrial Cancer Model with Retroperitoneal Lymphadenopathy Made From In Vivo Propagated and Cultured VX2 Cells.</a> J. Vis. Exp. (2019)
This video demonstrates the generation of a rabbit model of endometrial cancer by injecting a highly metastatic VX2 cell suspension into the sutured uterine horns of the rabbit. This leads to generation of primary endometrial cancer with retroperitoneal lymph node metastases.

<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/20371/20371_Figure1.jpg" /> 
Figure 1: Uterine suturing.&#160;Black arrow = sutures, red arrow = uterine horns.

Modeling Metastatic Endometrial Cancer  Model: A Technique to Generate Endometrial Cancer with Lymph Node Metastasis in Rabbit Models

Source: Lauren Philp et al. An Orthotopic Endometrial Cancer Model with Retroperitoneal Lymphadenopathy Made From In Vivo Propagated and Cultured VX2 ...

To begin, prep an anesthetized female rabbit in the dorsal position. Make a surgical incision through its abdominal wall to expose the abdominal cavity. Locate the uterine horns. These tubular structures are analogous to two distinct uteri extending from the fallopian tubes and opening up into two separate cervices on both sides. Surgically tie up the uterine horns on either side at a site closer to the cervices to isolate the distal uterine horn regions.
Now, inject a highly metastasizing desired tumor cell suspension into the myometrium or the outer muscular layer of the uterus at a point between the suture site and the cervix. Place the uterine horns back into the abdomen. Close the incision and allow the rabbit to recover. Within days of injection, a tumor develops at the site of injection. The cells from this tumor, with high metastatic potential, may travel through the lymphatic system and extravasate into lymph nodes, resulting in lymph node metastases.

modeling-metastatic-endometrial-cancer-model-technique-to-generate

Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Cancer Research

Gynecologic Cancer

Animal model studies

1.0K Views. Source: Lauren Philp et al. An Orthotopic Endometrial Cancer Model with Retroperitoneal Lymphadenopathy Made From In Vivo Propagated and Cultured VX2 Cells. J. Vis. Exp. (2019) This video demonstrates the generation of a rabbit model of endometrial cancer by injecting a highly metastatic VX2 cell suspension into the sutured uterine horns of the rabbit. This leads to generation of primary endometrial cancer with retroperitoneal lymph node metastases. 

Video: Modeling Metastatic Endometrial Cancer  Model: A Technique to Generate Endometrial Cancer with Lymph Node Metastasis in Rabbit Models - Concept

Development and Angiographic Use of the Rabbit VX2 Model for Liver Cancer

The rabbit VX2 tumor is an animal model commonly utilized for translational research regarding hepatocellular carcinoma (HCC) in the field of Interventional Radiology. This model employs an anaplastic squamous cell carcinoma that is easily and reliably propagated in the skeletal muscle of donor rabbits for eventual harvest and allograft implantation into the liver of na&#239;ve recipients. This tumor graft rapidly grows within the liver of recipient rabbits into an angiographically identifiable tumor characterized by a necrotic core surrounded by a viable hypervascular capsule. The physical size of the rabbit anatomy is sufficient to facilitate vascular instrumentation allowing for the application and testing of various interventional techniques. Despite these benefits, there exists a paucity of technical resources to act as a concrete reference for researchers working with the model. Herein, we present a comprehensive visual outline for the technical aspects of development, growth, propagation, and angiographic utilization of the rabbit VX2 tumor model for use by novice and experienced researchers alike.

The goal of this article is to provide a primer for the development and use of the VX2 carcinoma rabbit model for liver cancer.

The rabbit VX2 tumor is an animal model commonly utilized for translational research regarding hepatocellular carcinoma (HCC) in the field of ...

JoVE Journal

Two Methods for Establishing Primary Human Endometrial Stromal Cells from Hysterectomy Specimens

Many efforts have been devoted to establish in vitro cell culture systems. These systems are designed to model a vast number of in vivo processes. Cell culture systems arising from human endometrial samples are no exception. Applications range from normal cyclic physiological processes to endometrial pathologies such as gynecological cancers, infectious diseases, and reproductive deficiencies. Here, we provide two methods for establishing primary endometrial stromal cells from surgically resected endometrial hysterectomy specimens. The first method is referred to as &#8220;the scraping method&#8221; and incorporates mechanical scraping using surgical or razor blades whereas the second method is termed &#8220;the trypsin method.&#8221; This latter method uses the enzymatic activity of trypsin to promote the separation of cells and primary cell outgrowth. We illustrate step-by-step methodology through digital images and microscopy. We also provide examples for validating endometrial stromal cell lines via quantitative real time polymerase chain reactions (qPCR) and immunofluorescence (IF).

Establishing primary endometrial stromal cell culture systems from hysterectomy specimens is a valuable biological technique and a crucial step prior to pursuing a vast array of research aims. Here, we describe two methods used to establish stromal cultures from surgically resected endometrial tissues of human patients.&#160;

Many efforts have been devoted to establish in vitro cell culture systems. These systems are designed to model a vast number of in vivo processes. Cell ...

Medicine

Identification of Neutrophil Extracellular Traps in Paraffin-Embedded Feline Arterial Thrombi using Immunofluorescence Microscopy

Neutrophil extracellular traps (NETs), composed of cell-free DNA (cfDNA) and proteins like histones and neutrophil elastase (NE), are released by neutrophils in response to systemic inflammation or pathogens. Although NETs have previously been shown to augment clot formation and inhibit fibrinolysis in humans and dogs, the role of NETs in cats with cardiogenic arterial thromboembolism (CATE), a life-threatening complication secondary to hypertrophic cardiomyopathy, is unknown. A standardized method to identify and quantify NETs in cardiogenic arterial thrombi in cats will advance our understanding of their pathological role in CATE. Here, we describe a technique to identify NETs in formaldehyde-fixed and paraffin-embedded thrombi within the aortic bifurcation, extracted during necropsy. Following deparaffinization with xylene, aortic sections underwent indirect heat-induced antigen retrieval. Sections were then blocked, permeabilized, and ex vivo NETs were identified by colocalization of cell-free DNA (cfDNA), citrullinated histone H3 (citH3), and neutrophil elastase (NE) using immunofluorescence microscopy. To optimize the immunodetection of NETs in thrombi, autofluorescence of tissue elements was limited by using an autofluorescence quenching process prior to microscopy. This technique could be a useful tool to study NETs and thrombosis in other species and offers new insights into the pathophysiology of this complex condition.

We describe a method to identify neutrophil extracellular traps (NETs) in formaldehyde-fixed and paraffin-embedded feline cardiogenic arterial thrombi using heat-induced antigen retrieval and a double immunolabeling protocol.

Modeling Metastatic Endometrial Cancer Model: A Technique to Generate Endometrial Cancer with Lymph Node Metastasis in Rabbit Models

Transcript

Modeling Metastatic Endometrial Cancer Model: A Technique to Generate Endometrial Cancer with Lymph Node Metastasis in Rabbit Models

Development and Angiographic Use of the Rabbit VX2 Model for Liver Cancer

Two Methods for Establishing Primary Human Endometrial Stromal Cells from Hysterectomy Specimens

Identification of Neutrophil Extracellular Traps in Paraffin-Embedded Feline Arterial Thrombi using Immunofluorescence Microscopy