Quantifying Inferior Vena Cava Compliance and Distensibility in an In Vivo Ovine Model Using 3D Angiography

Abstract

Synthetic vascular grafts overcome some challenges of allografts, autografts, and xenografts but are often more rigid and less compliant than the native vessel into which they are implanted. Compliance matching with the native vessel is emerging as a key property for graft success. The current gold standard for assessing vessel compliance involves the vessel's excision and ex vivo biaxial mechanical testing. We developed an in vivo method to assess venous compliance and distensibility that better reflects natural physiology and takes into consideration the impact of a pressure change caused by flowing blood and by any morphologic changes present.

This method is designed as a survival procedure, facilitating longitudinal studies while potentially reducing the need for animal use. Our method involves injecting a 20 mL/kg saline bolus into the venous vasculature, followed by the acquisition of pre and post bolus 3D angiograms to observe alterations induced by the bolus, concurrently with intravascular pressure measurements in target regions. We are then able to measure the circumference and the cross-sectional area of the vessel pre and post bolus.

With these data and the intravascular pressure, we are able to calculate the compliance and distensibility with specific equations. This method was used to compare the inferior vena cava's compliance and distensibility in native unoperated sheep to the conduit of sheep implanted with a long-term expanded polytetrafluorethylene (PTFE) graft. The native vessel was found to be more compliant and distensible than the PTFE graft at all measured locations. We conclude that this method safely provides in vivo measurements of vein compliance and distensibility.

Introduction

Patients with critical cardiac anomalies require reconstructive surgery. Most reconstructive operations require the use of prosthetic materials, including vascular grafts. Potential conduits to bridge this space include synthetic or biologic materials. Initially, homografts were used as the Fontan conduit but have since been abandoned due to a high incidence of calcification and acute phase incidents¹. Currently, synthetic vascular grafts derived from inorganic polymers are used. There remains a challenge that these grafts are less compliant than the native vessel into which they are implanted and have long-term complications, such as stenosis, occlusion, and calcification^1,2,3,4,5.

The structure of synthetic vascular grafts lends itself to mechanical tensile strength, leading to their invariably lower compliance compared to native tissue². Vascular compliance, which defines the vessel's change in volume over a change in pressure, serves as an indicator of a vessel's responsiveness to mechanical loads. The difference between the graft material and native vessel properties creates a compliance mismatch, which has been demonstrated to disrupt blood flow patterns, resulting in areas of recirculation and flow separation^2,6,7,8,9. This phenomenon alters the shear stress on the endothelial wall and induces intimal hyperplasia^2,7,8,9. Such responses can lead to graft-related complications, necessitating graft replacement or re-intervention⁶.

As vascular compliance assumes a key role in determining graft outcomes, the accurate measurement of this property is essential. The current gold standard for measuring vascular compliance is ex vivo tubular biaxial mechanical testing. This method involves excising a graft or vessel of interest, connecting it to latex tubes, and pressurizing it to assess circumferential stress-stretch behavior across various pressures. Compliance is determined by comparing the pressure with a measurement of the inner diameter¹⁰. However, ex vivo methods have some disadvantages. When evaluating the functionality of implanted grafts using the ex vivo method, sacrificing the animals and explanting the grafts are necessary, making it impossible to conduct prolonged examinations. Therefore, we have developed an in vivo compliance measurement protocol.

Our group focuses on developing tissue-engineered vascular grafts (TEVGs) for use in the Fontan surgery to ameliorate the congenital heart defect hypoplastic left heart syndrome (HLHS). Recent developments in the field of congenital heart surgery have improved postoperative outcomes, leading to longer life expectancies. This makes the long-term properties and success of the implanted vascular conduit increasingly crucial. Currently, no animal model of HLHS exists so we evaluate our grafts in an accelerated large animal inferior vena cava (IVC) interposition graft model. While this model does not attempt to create the flow of the Fontan circulation, it effectively recapitulates the unique hemodynamic conditions. Our recent use of this in vivo protocol demonstrated significant differences in graft compliance between our TEVG and conventional expanded polytetrafluoroethylene (PTFE) grafts¹¹. As this previous study did not focus on methodology, we have conducted additional experiments detailing this novel in vivo method.

We implanted the synthetic graft currently serving as the standard of care, comprised of expanded polytetrafluoroethylene (PTFE), in Dorset sheep study animals and compared it to the native IVC in surgically naïve control animals. This protocol was performed on the PTFE group 5-7 years postimplantation of a PTFE conduit and unoperated control animals of varying ages. Thus, in subsequent sections describing the protocol and representative results, we will occasionally refer to the region of interest as, for example, the middle of the graft (midgraft) region of the IVC interposition graft.

This protocol allows us to analyze the in vivo compliance of the PTFE conduit, known to be non-compliant at a long-term time point, with the native vein. We chose to compare the clinical standard material, PTFE, with the native unoperated vein. We selected a long-term time point because the PTFE conduit is known to remain non-compliant and is prone to calcify, further reducing its compliance¹¹. We opted to conduct all comparisons in vivo as systemic hemodynamic changes are accurately reflected in measurements obtained through in vivo methods. From this comparison, we found that this protocol is able to confirm the non-compliance of PTFE and obtain measurements of in vivo venous compliance in a safe and reproducible manner. This method has been successfully implemented in a published study to demonstrate statistically significant differences between PTFE conduits and tissue-engineered vascular grafts (TEVGs) in vivo¹¹.

The overall goal of this protocol is to calculate compliance and distensibility of the thoracic IVC in an ovine large animal model using in vivo measurements from a survival procedure. To this end, we visualized and measured the changes in circumference and cross-sectional area of thoracic IVC to a fluid bolus. We simultaneously measured the intravascular change in pressure and used these measurements to calculate compliance and distensibility. Using 3D angiography imaging allows us multiple advantages, including the ability to adjust the view of the image post capture to ensure our measurements are taken from a cross-section of the vein, as well as allow us to measure multiple locations along the vessel. The three areas of interest in this study were the midgraft region, as well as the two adjacent anastomosis sites of the PTFE graft, and the comparable areas in the native IVC. By conducting experiments in vivo, there are advantages in evaluating the functionality of grafts within the actual flow of blood and surrounded by tissues and organs. The measurements obtained through this method are believed to reflect the actual functionality of the grafts in a living organism.

The protocol is divided into six main sections including preprocedure preparation of the sheep, catheterization, collection of baseline pre bolus data, collection of study data, animal recovery, and data analysis. In the animal preparation section, we discuss sedation, initiating anesthesia, and the placement of monitoring equipment used during the catheterization procedure. In the second section, we explain the process of placing the two catheter sheaths needed for data acquisition. For this protocol, both sheaths are placed in the right internal jugular vein (IJV) to allow two multitrack catheters to be introduced into the vessel. One will be positioned in the region of interest to record the change in pressure, and the other will be placed lower in the vein for contrast injection. Once the catheters are placed, a baseline pre bolus 3D angiograph is taken for comparison. Study data collection begins with preparing the saline bolus in a pressurized bag system for administration, providing the saline bolus with recording intravascular pressures, and taking the post bolus 3D angiograph. We then describe the process to facilitate the recovery of the sheep after the protocol. Lastly, we discuss the method to obtain the proper images and cross-sectional measurements for analysis and statistical comparison.

Protocol

The study protocol was approved by the Institutional Animal Care and Use Committee of Nationwide Children's Hospital Abigail Wexner Research Institute (AR22-0004). All animals received humane care in compliance with the Guide for the Care and Use of Laboratory Animals, published by the National Institutes of Health.

1. Animal preparation

1. Have a veterinary team evaluate the sheep 1 week prior to catheterization, including a physical exam and analysis of vital signs, to ensure the animal can safely receive anesthesia.
2. Fast the animal overnight, or up to 12 h prior to the procedure, to limit the risk of aspiration of stomach contents upon induction of anesthesia.
3. Press the on button on the control panel to turn on the C-arm and 3D angiography system (Figure 1A). Wait until the system is fully loaded.
   NOTE: Ensure the fluoroscopy is paused until ready to image and all personnel are wearing protective lead.
4. Prep heparinized saline to use in the procedure by adding 1 mL of heparin (1,000 USP units/mL concentration) to 1,000 mL of 0.9% saline.
5. Shave the left side of the neck and scrub with alcohol. Administer sedation by injecting a combination of ketamine (4 mg/kg), butorphanol (0.1 mg/kg), and diazepam (0.5 mg/kg) into the left jugular vein.
6. Place the sedated sheep on a hospital bed and position it in sternal recumbency for intubation. Intubate with a 9-14 mm Endotracheal (ET) tube, based on the sheep's size, by depressing the tongue and epiglottis with a laryngoscope and inserting the ET tube into the trachea.
7. Position the sheep in a right lateral position. Attach the ET tube to a ventilator and mechanically ventilate with 100% Oxygen at 1-3 L/min.
8. Maintain anesthesia with 1-3% inhaled isoflurane. Set the respiratory rate at 15-30 breaths/min and end tidal volume at 8-10 mL/kg.
9. Place standard monitoring equipment, including a blood pressure cuff on the right front leg, an ear clip to monitor oxygen saturation on the right ear, a temperature probe into the esophagus, and an end tidal CO₂ monitor on the ET tube. Shave the wool from the caudal aspect of each hoof between the dewclaws and heel. Place electrocardiogram (ECG) nodes and secure the ECG nodes with tape.
10. Lubricate both eyes by applying ophthalmic ointment and insert an orogastric tube to ensure gas and food evacuation.
11. Establish an IV line in the left IJV to allow for the administration of propofol constant-rate infusion (CRI) (20-40 mg∙kg^-1∙h^-1), maintenance fluids (10 mL∙kg^-1∙h^-1), and the saline bolus.
12. Position the sheep in left lateral recumbency. Shave the right side of the neck to gain access to the catheterization site (Figure 2A). Swab the area with chlorhexidine scrub and alcohol.
13. Disconnect from the monitoring equipment and ventilator and move the sheep to the catheterization laboratory table. Again, situate the animal in a left lateral recumbent position (Figure 3A).
14. Reconnect to the ventilator and monitoring equipment (ECG leads, temperature probe, blood pressure cuff, pulse oximeter).
15. Maintain anesthesia during the procedure by administering inhaled isoflurane 1-3% with 100% O₂, and/or propofol CRI (20-40 mg∙kg^-1∙h^-1).
    NOTE: Assess the plane of anesthesia by gauging animal movement, response to painful stimuli, respiratory rate, pulse rate, and blood pressure. Make adjustments in sedation as appropriate, such as by using a 5-10 mL bolus of propofol to induce a deeper plane of anesthesia.
16. Measure the sheep's width at the area of the heart using large calipers. Divide the width by 2 to set the pressure transducer.
17. Aseptically clean the surgical site and drape it in a sterile fashion (Figure 2B,C).

2. Catheterization

1. Move the C-Arm from the parked position to the sheep's chest and raise the table as needed. Press buttons 7 and 3 on the control panel, and then hold the Start button to use the preprogrammed settings to autoposition the table and the C-Arm on the left side of the table (Figure 1A).
2. Access the right IJV using a 21 G micro puncture needle and 10 cc Luer Slip syringe; access the IJV in a cranial/caudal direction through the skin while pulling back on the syringe plunger. Ensure that blood is aspirated to confirm that the needle is in the vessel (Figure 2A, B).
3. Carefully disconnect the syringe while holding the needle steady.
4. Insert a 0.018 inch Stainless Steel (SS) Wire Guide through the needle into the vessel approximately halfway. Remove the needle from over the SS wire.
5. Place a 5-French (Fr) dilator over the SS wire and into the vessel. Remove the inner piece of the dilator and SS wire. Feed a 0.038 inch guidewire through the dilator into the vessel about halfway and remove the dilator.
6. Use an 11-blade scalpel to cut the skin above the vein where the wire enters. Feed a 9-Fr sheath over the guidewire and into the vessel. Remove the inner sheath section and guidewire.
7. Confirm proper sheath placement by aspirating blood and then flushing the sheath with heparinized saline.
8. Repeat steps 2.2-2.7 so that there are two 9-Fr sheaths in the right IJV.
9. Administer 150 U/kg of heparin through the IV to prevent clotting.
10. Use the foot pedal to start fluoroscopy (Figure 1B). Insert a Judkins Right (JR) catheter through the sheath, following the thoracic IVC across the diaphragm into the abdominal IVC.
11. Thread a Rosen wire through the JR catheter until it reaches the abdominal IVC and the tip emerges from the JR catheter. While holding the Rosen wire in place, gently remove the JR catheter.
12. Repeat steps 2.10-2.11 with the second sheath.
13. Thread a 7-Fr Multitrack catheter over each Rosen wire.
14. Under fluoroscopy guidance, place one multitrack angiographic catheter in the abdominal IVC for contrast injection.
15. Using fluoroscopy, place another multitrack angiographic catheter into the specific region of interest (e.g., the center of the graft) for pressure measurement (Figure 2C).

Figure 1: Control panel. (A) 3D angiography system control panel (B) Fluoroscopy foot pedals Please click here to view a larger version of this figure.

Figure 2: Animal catheterization. (A) The key surgical site, prepped for catheterization. (B) Technique to visualize the right internal jugular vein (black arrows). (C) Two multitrack angiographic catheters are placed through the right internal jugular vein (blue arrow: pressure measurement in the graft; red arrow: contrast injection into abdominal IVC; white arrow: stiff wires). Abbreviation: IVC = inferior vena cava. Please click here to view a larger version of this figure.

3. Gathering the predata

1. Connect the multitrack centered in the region of interest to the pressure transducer using a three-way stopcock. With the stopcock open to the multitrack, pull back with a 10 mL syringe until air bubbles are removed and blood is seen.
2. With the 10 mL syringe flipped upside-down, return the blood to the sheep with care not to push any air back into the multitrack. Flush the multitrack with heparinized saline.
3. Flip the off position of the stopcock to the syringe so that the pressure transducer and the multitrack are open to each other.
4. Prepare the contrast injector by adding a contrast agent. The minimal volume for a 3D angiogram is 60 mL and the total contrast cannot exceed 5 mL/kg or 250 mL.
5. Connect the contrast injector to the multitrack centered in the abdominal IVC. Using the contrast injector, turn the knob counterclockwise slowly to pull air bubbles from the multitrack until blood is seen. Turn the knob clockwise to push contrast forward slowly into the multitrack.
6. Use fluoroscopy to confirm when the contrast has reached the tip of the multitrack.
7. Take the total contrast used for the 3D angiogram and divide by 5 to get mL/s. Set the rate rise to 0 and 600 psi.
8. Set the C-arm to the preprogrammed mode by clicking the Program button on the right top of the screen and the 3D DSA 110 8'' button (3D Angiography with an SID of 110 cm and Field View at 8 inches).
9. Move all objects and people away from the front or sides of the table. Initiate the C-Arm program by clicking the button numbered 3 on the control panel. Position the target region (e.g., midgraft) at the center of the x-y plane (Figure 3A-C).
10. Progress to the second program by clicking the button numbered 4 on the control panel. Adjust the height of the table accordingly to center the region of interest.
11. Press the button numbered 5 and take the test image.
12. Pretest the C-Arm's range of motion by clicking the Confirm Conditions button | Start (Figure 3D,E).
13. Ask the anesthesiologist to hold ventilation and take a 3D rotational angiogram with contrast injection by initiating the program with the middle acquisition pedal. Concurrently measure and record the mean pressure of the target region.

Figure 3: C-arm positioning and range of motion. (A) Sheep positioning for the beginning of the procedure (B) First position for 3D angiogram program (C) C-arm moved in the xy-axis (D) C-arm moved in the z-axis (E) C-arm completing test-spin with full range of motion. Please click here to view a larger version of this figure.

4. Administering the saline bolus and collecting data

1. Prep 20 mL/kg of 0.9% saline.
2. To prepare the bolus pressurized bag, add a 1,000 mL bag of 0.9% saline into a pressurized bag unit. Use a second unit if necessary to achieve the total volume to be administered.
3. Squeeze the inflation bulb until the pressure gauge rises into the green zone, immediately before the red line (pressure 250-300 mmHg). Flush saline through the line and remove air bubbles.
4. Connect the pressurized saline bag to a 9-Fr sheath and maintain 250-300 mmHg to maintain a constant speed of the bolus. Allow it to flow until the sheep has received a bolus equivalent to 20 mL/kg or the mean pressure reaches 15 mmHg.
5. While the bolus is flowing, record intravascular pressures of the target region every minute.
6. Have the C-Arm ready for a second 3D rotational angiography by repeating steps 3.9-3.12. As soon as the bolus ends, before the pressure begins to drop, take the second 3D angiography and concurrent intravascular pressure measurement by initiating as described in step 3.13.

5. Recovery

1. Following imaging completion, place the C-Arm back in the preprogrammed parked position by entering number 77 and holding the Start button until the C-Arm positions itself.
2. Remove the multitrack angiography catheters and Rosen wires.
3. Remove both sheaths while applying direct pressure over insertion sites with a hemostasis patch for at least 7 min to stop bleeding.
4. Wrap a sterile roll of gauze around patches and neck such that the wrap is secure to maintain pressure, but not too tight to risk cutting off circulation or preventing breathing.
5. Turn off anesthetics (isoflurane and/or propofol CRI).
6. Maintain sheep on the ventilator with 100% O₂ until they consistently breathe on their own.
   NOTE: Signs that the sheep is waking up include movement, blinking, response to painful stimuli, jaw tone or attempts to chew, and breathing without assistance from a ventilator.
7. Once the sheep is able to breathe on their own, extubate (remove the ET tube) and remove the orogastric tube.
8. Remove all monitoring equipment and transfer the sheep to a hospital bed. Move it back to the housing room.
9. Assist the sheep with staying in sternal recumbency or when attempting to stand until the sheep is able to keep balance on their own. Prevent them from running into walls.
10. Once they appear awake enough, give small amounts of hay or grain.

6. Analysis of data

1. Export the original raw 3D angiography data from the angiography imaging software in DICOM file format.
2. Launch the DICOM viewer software. Drag and drop the 3D angiography file into the viewer to open (Figure 4A).
3. Within the DICOM viewer software, select the 3D MPR (Multi-Planar Reconstruction) tool to generate a 3D reconstructed view of the angiography data. This will present three distinct 2D views from three different angles: axial, sagittal (Figure 4B), and coronal (Figure 4C) planes.
4. Adjust the placement and orientation of the target region in the sagittal and coronal planes to achieve the desired vertical position by placing the target region at the center and rotating the direction of the reference lines on each plane with a hand tool (Figure 4D).
5. Utilize the pencil tool within the DICOM viewer to outline the vessel wall in the axial view of the target region (Figure 4E). The software automatically calculates and shows both the area and perimeter (Circumference) of the region in the middle of the axial view.
6. Utilize equation (1) to calculate compliance, where A is the cross-sectional area (cm²) and P is the pressure (mmHg):
         (1)
7. Employ equation (2) to calculate distensibility where C is the circumference (cm) and P is the pressure (mmHg):
      (​2)

Figure 4: Data analysis in DICOM viewer. (A) Raw data of 3D angiogram loaded into DICOM viewer. (B) The sagittal section of the graft. (C) The coronal section. (D) A true cross-section is visualized after adjusting the angle on the sagittal and coronal sections. (E) The pencil tool is used to outline the target vessel to make circumference and cross-sectional area measurements. Please click here to view a larger version of this figure.

Representative Results

We have successfully performed this procedure with over 25 sheep. Importantly, there were no instances of morbidity and mortality related to this procedure. All the sheep exhibited uncomplicated recoveries. These representative results were taken from three sheep implanted with PTFE grafts and three unoperated native sheep. Figure 5 provides the intravascular pressure measurements taken from both groups of study animals during the protocol. These values are important for the compliance and distensibility calculations, but also to demonstrate the safety of this protocol as we maintain the change in intravascular pressure below 15 mmHg.

Figure 5: Intravascular pressures for compliance and distensibility equations. (A) Graphical representation of the absolute intravascular pressure measurements taken at the midgraft area, or corresponding area of the native vessel, during the course of bolus administration. (B) Summary table of intravascular pressure changes for each study animal. These values were used in compliance and distensibility calculations. Please click here to view a larger version of this figure.

Figure 6A,B are representative images of the DICOM viewer measurements in the native and PTFE groups. The figure demonstrates the change in circumference seen in the native sheep in response to the bolus, which is not seen in the PTFE group. From the measurements, we calculated the compliance and distensibility, which are presented in Figure 6C. These values were then graphed and statistically analyzed. The comparison of the compliance and distensibility between native sheep and PTFE sheep demonstrates that native sheep vessels are more compliant and distensible than PTFE grafts. The observed differences all trended towards statistical significance, with only the distal compliance and proximal compliance comparisons being statistically significant (p < 0.05).

Figure 6: Representative results. (A) Representative images of the circumference of the native vessel measured in the DICOM viewer pre and post bolus. A circumferential outline (green) was traced in the DICOM viewer. Circumference and cross-sectional area values (boxed green text) were then automatically generated by the DICOM viewer. (B) Representative images of the circumference of the PTFE graft measured in the DICOM viewer pre and post bolus. A circumferential outline (green) was traced in the DICOM viewer. Circumference and cross-sectional area values (boxed green text) were then automatically generated by the DICOM viewer. (C) Table of calculated compliance and distensibility for each study animal at each vessel location. Negative compliance and distensibility values were adjusted to zero. (D) Representation of compliance and distensibility data. Data normality was tested prior to all statistical comparisons. Compliance and distensibility values were analyzed with an unpaired Student's t-test, where Welch's correction was applied for all measurements except for distal compliance. Distal and proximal compliance values were significantly higher in the native group compared to the PTFE group (distal: p = 0.0485, proximal: p = 0.0247). The midgraft compliance was not statistically significant between groups, yet native midgraft compliance was always higher than the PTFE. Abbreviation: PTFE = polytetrafluoroethylene. Please click here to view a larger version of this figure.

Discussion

Compliance and distensibility are key properties for blood vessel function, serving as indicators of potential complications and interventions. Precisely quantifying and comparing changes in these parameters is important to assess graft efficacy. Our in vivo method overcomes the limitations of ex vivo analysis and maintains comparable results. Comparing our in vivo data to the ex vivo data presented by Blum et al., both methods demonstrate marked differences between the synthetic graft material of interest and the native vein¹⁰. Our statistical significance was limited by the low sample size. Despite their focus on a different graft conduit, we find the protocols to be scientifically comparable in regard to data outputs, with the in vivo method being preferable for the reasons previously mentioned.

Several critical steps in this procedure demand careful attention, particularly in the catheter insertion and bolus administration. If accessing the vein proves challenging, adjusting the sheep's head angle or using positioning aids can be beneficial. Placing the JR catheter in the precise area of interest can be facilitated by using a glide wire. Another critical step may occur when switching saline bags due to a large bolus requirement. A quick switch is essential to avoid significant intravascular pressure fluctuations. To avoid this issue, we prepared both saline bags to match the necessary volume exactly and connected both to the sheath with a stopcock. Prior to the bolus initiation, we pressurized both bags, reducing the time required for the flow transition from one bag to the other. It is crucial to maintain a time-efficient approach as extended anesthesia periods can negatively impact sheep recovery.

It is critical to test the range of the C-Arm before initiating the protocol. While testing the C-Arm, ensure that it centers the region of interest accurately. Repositioning the C-Arm properly will guarantee full visualization of the area of interest in the final image. Timing is critical to image the bolus effectively. Initiate the imaging sequence as soon as the pressurized bag is emptied. Ensuring that the test spin of the C-Arm is done prior to administering fluid can help in catching this critical moment.

Of note, 2D angiography imaging is not a suitable substitute for 3D angiography. The compliance and distensibility equations require precise cross-sectional area and circumference values, which 2D imaging cannot provide due to natural vessel angles and variations in morphology. Intravascular ultrasound, though considered, is technically challenging and lacks post-capture adjustability.

This method is currently applicable to venous vasculature and will require modification before being applied to arterial circulation. Potential applications of this method include comparing transplanted or graft materials in the venous system, as well as assessing compliance changes associated with aging longitudinally. Before the administration of a fluid bolus, the current method using 3D angiography did not eliminate the effect of IVC pulsatility from the heartbeat.

Materials

Name	Company	Catalog Number	Comments
0.035" x 260 cm Rosen Curved Wire Guide	Cook Medical	G01253	Guide for holding placement swapping caths (Multi-track, IVUS, etc)
0.035"x 150 cm Glidewire	Terumo	GR3507	Guide for JR cath
0.9% Sodium Chloride Saline	Baxter Healthcare Corporation	NCH pharmacy	For diluting norepinpherine, pressure monitoring
10.0 Endotracheal tube	Coviden	86117	To secure airway
16 G IV catheter	BD	382259	To administer fluids and anesthetic drugs
22 G IV catheter	BD	381423	For invasive blood pressure
5Fr x .35" JR2.5	Cook Medical	G05035	Guide for rosen wire
70% isopropyl alcohol	Aspen Vet	11795782	Topical cleaning solution
7Fr x 100 cm Multi-track	B. Braun	615001	Collecting pressure, Administering contrast to specific intravascular location
9Fr Introducer sheath	Terumo	RSS901	Access catheter through skin into vessel for wires to pass through
ACT cartridge	Abbot Diagnostics	03P86-25	Activated Clotting Time
Angiographic syringe w/ filling spike	Guerbet	900103S	For contrast injector
Bag decanter	Advance Medical Designs, LLC	10-102	Punctures saline bag to pour and fill sterile bowl with saline
Butorphanol	Zoetis	NCH pharmacy	Sedation drug: Concentration 10 mg/mL, Dosage 0.1 mg/kg
Cath Research Pack	Cardinal Health	SAN33RTCH6	Cath pack with misc. supplies
Cetacaine	Cetylite	220	Topical anesthetic spray
Chloraprep	BD	930825	Topical cleaning solution
Chlorhexidine 2% solution	Vedco INC	VINV-CLOR-SOLN	Topical cleaning solution
Conform stretch bandage	Coviden	2232	Neck wrap to prevent bleeding
Connection tubing	Deroyal	77-301713	Connects t-port to fluid/drug lines
Diazepam	Hospira Pharmaceuticals	NCH pharmacy	Sedation drug: Concentration 5 mg/mL, Dosage 0.5 mg/kg
EKG monitoring dots	3M	2570
Fluid administration set	Alaris	2420-0007
Fluid warming set	Carefusion	50056
Hemcon Patch	Tricol Biomedical	1102	Patch for hemostasis
Heparin	Hospira, Inc	NCH pharmacy	Angicoagulant: 1,000 USP units/mL
Infinix-i INFX-8000C	Toshiba Medical Systems	2B308-124EN*E	Interventional angiography system
Invasive pressure transducer	Medline	23DBB538	For invasive blood pressure
Isoflurane	Baxter Healthcare Corporation	NCH pharmacy	Anesthetic used in prep room
Ketamine	Hospira Pharmaceuticals	NCH pharmacy	Sedation drug: Concentration 100 mg/mL, Dosage 4 mg/kg
Lubricating Jelly	MedLine	MDS0322273Z	ET tube lubricant
Micropuncture Introducer Set	Cook Medical	G47945	Access through skin into vessel
Needle & syringes	Cardinal Health	309604	For sedation
Norepinpherine Bitartrate Injection, USP	Baxter Healthcare Corporation	NCH pharmacy	1 mg/mL
Optiray 320	Liebel-Flarsheim Company, LLC	NCH pharmacy	Contrast
Optixcare	Aventix	OPX-4252	Corneal lubricant
OsiriX MD	Pixmeo SARL	-	DICOM Viewer and Analysis software
Pressure infusor bag	Carefusion	64-10029	To maintain invasive blood pressure
Propofol	Fresenius Kabi	NCH pharmacy	Anesthetic drug: Concentration 10 mg/mL, Dosage 20-45 mg·kg-1·h-1
Silk suture 3-0	Ethicon	C013D	To secure IV catheter
SoftCarry Stretcher	Four Flags Over Aspen	SSTR-4
Stomach tube	Jorgensen Lab, INC	J0348R	To release gastric juices and gas and prevent bloat
T-port	Medline	DYNDTN0001	Connects to IV catheter
Urine drainage bag	Coviden	3512	Connects to stomach tube to collect gastric juices
Warming blanket	Jorgensen Lab, INC	J1034B