Name: quantifying inferior vena cava compliance and distensibility in an in vivo ovine model using 3d angiography
Uploaded: 2024-04-26T14:50:00.000+00:00
Duration: 7 min 53 s
Description: Read this Scientific Article Protocol about Author Spotlight: Improved Imaging for Neovascular Development in Congenital Heart Disease Research at JoVE.com

This work was supported by R01 HL163065 and W81XWH1810518. We extend our appreciation to the dedicated staff at the Animal Research Core. We also wish to express our gratitude to Carmen Arsuaga for her invaluable expertise and vigilant care throughout the study.

The study protocol was approved by the Institutional Animal Care and Use Committee of Nationwide Children&#39;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
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#39;s size, by depressing the tongue and epiglottis with a laryngoscope and inserting the ET tube into the trachea.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Lubricate both eyes by applying ophthalmic ointment and insert an orogastric tube to ensure gas and food evacuation.</li>
	<li>Establish an IV line in the left IJV to allow for the administration of propofol constant-rate infusion (CRI) (20-40 mg&#8729;kg-1&#8729;h-1), maintenance fluids (10 mL&#8729;kg-1&#8729;h-1), and the saline bolus.</li>
	<li>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 at least 2 alcohol scrubs and 2 chlorhexidine scrubs.</li>
	<li>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).</li>
	<li>Reconnect to the ventilator and monitoring equipment (ECG leads, temperature probe, blood pressure cuff, pulse oximeter).</li>
	<li>Maintain anesthesia during the procedure by administering inhaled isoflurane 1-3% with 100% O2, and/or propofol CRI (20-40 mg&#8729;kg-1&#8729;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.</li>
	<li>Measure the sheep&#39;s width at the area of the heart using large calipers. Divide the width by 2 to set the pressure transducer.</li>
	<li>Aseptically clean the surgical site and drape it in a sterile fashion (Figure 2B,C).</li>
</ol>
2. Catheterization
<ol>
	<li>Move the C-Arm from the parked position to the sheep&#39;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).</li>
	<li>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).</li>
	<li>Carefully disconnect the syringe while holding the needle steady.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Confirm proper sheath placement by aspirating blood and then flushing the sheath with heparinized saline.</li>
	<li>Repeat steps 2.2-2.7 so that there are two 9-Fr sheaths in the right IJV.</li>
	<li>Administer 150 U/kg of heparin through the IV to prevent clotting.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Repeat steps 2.10-2.11 with the second sheath.</li>
	<li>Thread a 7-Fr Multitrack catheter over each Rosen wire.</li>
	<li>Under fluoroscopy guidance, place one multitrack angiographic catheter in the abdominal IVC for contrast injection.</li>
	<li>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).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66724/66724fig01.jpg" /> 
Figure 1: Control panel. (A) 3D angiography system control panel (B) Fluoroscopy foot pedals <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66724/66724fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66724/66724fig02v3.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66724/66724fig02largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Gathering the predata
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Flip the off position of the stopcock to the syringe so that the pressure transducer and the multitrack are open to each other.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Use fluoroscopy to confirm when the contrast has reached the tip of the multitrack.</li>
	<li>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.</li>
	<li>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&#39;&#39; button (3D Angiography with an SID of 110 cm and Field View at 8 inches).</li>
	<li>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).</li>
	<li>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.</li>
	<li>Press the button numbered 5 and take the test image.</li>
	<li>Pretest the C-Arm&#39;s range of motion by clicking the Confirm Conditions button | Start (Figure 3D,E).</li>
	<li>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.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66724/66724fig03.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66724/66724fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Administering the saline bolus and collecting data
<ol>
	<li>Prep 20 mL/kg of 0.9% saline.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>While the bolus is flowing, record intravascular pressures of the target region every minute.</li>
	<li>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.</li>
</ol>
5. Recovery
<ol>
	<li>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.</li>
	<li>Remove the multitrack angiography catheters and Rosen wires.</li>
	<li>Remove both sheaths while applying direct pressure over insertion sites with a hemostasis patch for at least 7 min to stop bleeding.</li>
	<li>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.</li>
	<li>Turn off anesthetics (isoflurane and/or propofol CRI).</li>
	<li>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.</li>
	<li>Once the sheep is able to breathe on their own, extubate (remove the ET tube) and remove the orogastric tube.</li>
	<li>Remove all monitoring equipment and transfer the sheep to a hospital bed. Move it back to the housing room.</li>
	<li>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. 
	NOTE: Monitor the animals closely during recovery. If any signs of pain or distress are observed, consult the veterinary team.</li>
	<li>Once they appear awake enough, give small amounts of hay or grain.</li>
</ol>
6. Analysis of data
<ol>
	<li>Export the original raw 3D angiography data from the angiography imaging software in DICOM file format.</li>
	<li>Launch the DICOM viewer software. Drag and drop the 3D angiography file into the viewer to open (Figure 4A).</li>
	<li>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.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Utilize equation (1) to calculate compliance, where A is the cross-sectional area (cm2) and P is the pressure (mmHg): 
	<img alt="Equation 1" src="/files/ftp_upload/66724/66724eq01.jpg" style="margin: auto;" />&#160; &#160; &#160; (1)</li>
	<li>Employ equation (2) to calculate distensibility where C is the circumference (cm) and P is the pressure (mmHg): 
	<img alt="Equation 2" src="/files/ftp_upload/66724/66724eq02.jpg" style="margin: auto;" />&#160; &#160;(&#8203;2)</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66724/66724fig04.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66724/66724fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

3D Angiography for Venous Compliance Assessment in Sheep

<ol>
	<li>Hagler, D. 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=Fate+of+the+Fontan+connection:+Mechanisms+of+stenosis+and+management.">Fate of the Fontan connection: Mechanisms of stenosis and management.</a> Congenit Heart Dis. 14 (4), 571-581 (2019).</li><li>Nezerati, R. M., Eifert, M. B., Dempsey, D. K., Cosgriff-Hernandez, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrospun+vascular+grafts+with+improved+compliance+matching+to+native+vessels.">Electrospun vascular grafts with improved compliance matching to native vessels.</a> J Biomed Mater Res B Appl Biomater. 103 (2), 313-323 (2015).</li><li>Bates, O., Semple, T., Krupickova, S., Bautisa-Rodriguez, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Case+report+of+a+Gore-Tex+TCPC+conduit+dissection+causing+severe+stenosis.">Case report of a Gore-Tex TCPC conduit dissection causing severe stenosis.</a> Eur Heart J Case Rep. 5 (11), 1-6 (2021).</li><li>Sathananthan, G., 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=Clinical+importance+of+Fontan+Circuit+thrombus+in+the+adult+population:+Significant+association+with+increased+risk+of+cardiovascular+events.">Clinical importance of Fontan Circuit thrombus in the adult population: Significant association with increased risk of cardiovascular events.</a> Can J Cardiol. 35 (12), 1807-1814 (2019).</li><li>Kumar, P., Bhatia, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Computed+tomography+in+the+evaluation+of+Fontan+Circulation.">Computed tomography in the evaluation of Fontan Circulation.</a> J Cardiovasc Imaging. 29 (2), 108-122 (2021).</li><li>Abbott, W. M., Megerman, J., Hasson, J. E., L'Italien, G., Warnock, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+compliance+mismatch+on+graft+patency.">Effect of compliance mismatch on graft patency.</a> J Vasc Surg. 5 (2), 376-382 (1987).</li><li>Weston, M. W., Rhee, K., Tarbell, 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=Compliance+and+diameter+mismatch+affect+the+wall+shear+rate+distribution+near+an+end-to-end+anastomosis.">Compliance and diameter mismatch affect the wall shear rate distribution near an end-to-end anastomosis.</a> J Biomech. 29 (2), 187-198 (1996).</li><li>Ballyk, P. D., Walsh, C., Butany, J., Ojha, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compliance+mismatch+may+promote+graft-artery+intimal+hyperplasia+by+altering+suture-line+stress.">Compliance mismatch may promote graft-artery intimal hyperplasia by altering suture-line stress.</a> J Biomech. 31 (3), 229-237 (1997).</li><li>Lemson, M. S., Tordoir, J. H. M., Daemen, M. J. A. P., Kitslaar, P. J. E. H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intimal+hyperplasia+in+vascular+grafts.">Intimal hyperplasia in vascular grafts.</a> Eur J Vasc Endovasc Surg. 19 (4), 336-350 (2000).</li><li>Blum, K. M., 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=Tissue+engineered+vascular+grafts+transform+into+autologous+neovessels+capable+of+native+function+and+growth.">Tissue engineered vascular grafts transform into autologous neovessels capable of native function and growth.</a> Commun Med. 2, 3 (2022).</li><li>Turner, M. E., 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=Tissue+engineered+vascular+grafts+are+resistant+to+the+formation+of+dystrophic+calcification.">Tissue engineered vascular grafts are resistant to the formation of dystrophic calcification.</a> Nat Commun. 15, 2187 (2024).</li></ol>

This study was sponsored in part by Gunze Ltd.

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.

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 calcification1,2,3,4,5.
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&#39;s change in volume over a change in pressure, serves as an indicator of a vessel&#39;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 separation2,6,7,8,9. This phenomenon alters the shear stress on the endothelial wall and induces intimal hyperplasia2,7,8,9. 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 na&#239;ve 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.

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

quantifying inferior vena cava compliance and distensibility in an in vivo ovine model using 3d angiography

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.

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.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66724/66724fig05.jpg" /> 
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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66724/66724fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
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 &#60; 0.05).
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66724/66724fig06.jpg" /> 
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&#39;s t-test, where Welch&#39;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. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66724/66724fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Read this Scientific Article Protocol about Author Spotlight: Improved Imaging for Neovascular Development in Congenital Heart Disease Research at JoVE.com

Author Spotlight: Improved Imaging for Neovascular Development in Congenital Heart Disease Research

This protocol allows for the in vivo quantification of venous compliance and distensibility using catheterization and 3D angiography as a survival procedure allowing for a variety of potential applications.

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

Synthetic vascular grafts overcome some challenges of allografts, autografts, and xenografts but are often more rigid and less compliant than the native ...

quantifying-inferior-vena-cava-compliance-distensibility-an-vivo

Nanyang Technological University

Research

JoVE Journal

Bioengineering

474 Views.  Nationwide Children's Hospital. We streven ernaar om de langetermijnresultaten voor patiënten met aangeboren hartaandoeningen te verbeteren met behulp van een regeneratieve geneeskundebenadering. Ons weefsel-gemanipuleerde vasculaire transplantaat ontwikkelt zich tot een neo-vat dat bestaat uit de eigen cellen van de patiënt. Ons doel is om vergelijkingen te maken tussen ons transplantaat, de native ader, en de klinische standaard polytetrafluorethyleen of PTFE.We hebben vastgestel...

Video: Auteur in de schijnwerpers: Verbeterde beeldvorming voor neovasculaire ontwikkeling in onderzoek naar aangeboren hartaandoeningen

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and behave by observing the surface of a tissue. This method provides a means to visualize the complex 3-dimensional vascular architecture of the lung.
To accomplish this, a catheter is inserted into the pulmonary artery and the vasculature is simultaneously flushed of blood and chemically dilated to limit resistance. Lungs are then inflated through the trachea at a standard pressure and the polymer compound is infused into the vascular bed at a standard flow rate. Once the entire arterial network is filled and allowed to cure, the lung vasculature may be visualized directly or imaged on a micro-CT (&#181;CT) scanner.
When performed successfully, one can appreciate the pulmonary arterial network in mice ranging from early postnatal ages to adults. Additionally, while demonstrated in the pulmonary arterial bed, this method can be applied to any vascular bed with optimized catheter placement and endpoints.

The aim of this technique is ex vivo visualization of pulmonary arterial networks of early postnatal and adult mice through lung inflation and injection of a radio-opaque polymer-based compound via the pulmonary artery. Potential applications for casted tissues are also discussed.

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and ...

Developmental Biology

Three-Dimensional Echocardiographic Method for the Visualization and Assessment of Specific Parameters of the Pulmonary Veins

The dimensions of the pulmonary veins are important parameters when planning pulmonary vein isolation (PVI), especially with the cryoballoon ablation technique. Acknowledging the dimensions and anatomical variations of the pulmonary veins (PVs) may improve the outcome of the intervention. Conventional 2D transoesophageal echocardiography can only provide limited data about the dimensions of the PVs; however, 3D echocardiography can further evaluate relevant diameters and areas of the PVs, as well as their spatial relationship to surrounding structures. In previous literature data, parameters influencing the success rate of PVI have already been identified. These are the left lateral ridge, the intervenous ridge, the ostial area of the PVs and the ovality index of the ostium. Proper imaging of the PVs by 3D echocardiography is a technically challenging method. One crucial step is the collection of images. Three individual transducer positions are necessary to visualize the important structures; these are the left lateral ridge, the ostium of the PVs and the intervenous ridge of the left and right PVs. Next, 3D images are acquired and saved as digital loops. These datasets are cropped, which result in the en face views displaying spatial relationships. This step can also be employed to determine the anatomical variations of the PVs. Finally, multiplanar reconstructions are created to measure each individual parameter of the PVs.
Optimal quality and orientation of the acquired images are paramount for the appropriate assessment of PV anatomy. In the present work, we examined the 3D visibility of the PVs and the suitability of the above method in 80 patients. The aim was to provide a detailed outline of the essential steps and potential pitfalls of PV visualization and assessment with 3D echocardiography.

The dimensions of the pulmonary veins (PV) are important parameters when planning pulmonary vein isolation. 2D transoesophageal echocardiography can only provide limited data about the PVs; however, 3D echocardiography can evaluate relevant diameters and areas of the PVs, as well as their spatial relationship to surrounding structures.

The dimensions of the pulmonary veins are important parameters when planning pulmonary vein isolation (PVI), especially with the cryoballoon ablation ...

Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ligation of the inferior vena cava (IVC) in the outflow of the fistula, mimicking central venous stenosis. Technical aspects of this model are introduced. The aorta and IVC are exposed, following an abdominal incision. The infra-renal aorta and IVC are dissected for proximal clamping, and the distal aorta is exposed for puncture. The IVC at the midpoint between the left renal vein and the aortic bifurcation is carefully dissected to place an 8-0 suture beneath the IVC. After clamping the aorta and IVC, an AVF is created by puncturing the infra-renal aorta through both walls into the IVC with a 25 G needle, followed by ligating a 22 G intra-venous (IV) catheter and IVC together. The catheter is then removed, creating a reproducible venous stenosis without occlusion. The aorta and IVC are unclamped after confirming primary hemostasis. This novel model of central vein stenosis is easy to perform, reproducible, and will facilitate studies on AVF failure.

An aortocaval fistula was created by puncturing the murine infra-renal aorta through both walls into the inferior vena cava and was followed by creation of a stenosis in its outflow via partial ligation of the inferior vena cava. This reproducible model can be used to study central venous stenosis.

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ...

Medicine

Measuring Cardiac Output in a Swine Model

Swine are frequently used in medical research given their similar cardiac physiology to that of humans. Measuring cardiac parameters such as stroke volume and cardiac output are essential in this type of research. Contrast ventriculography, thermodilution, and pressure-volume loop (PV-loop) catheters can be used to accurately obtain cardiac performance data depending on which resources and expertise are available. For this study,five Yorkshire swine were anesthetized and intubated. Central venous and arterial access was obtained to place the necessary measurement instruments.A temperature probe was placed in the aortic root. A cold saline bolus was delivered to the right atrium and temperature deflection curve was recorded. Integration of the area under the curve allowed for the calculation of the current cardiac output.A pigtail catheter was percutaneously placed in the left ventricle and 30 mL of iodinated contrast was power injected over 2 seconds. Digital subtraction angiography images were uploaded to volumetric analysis software to calculate the stroke volume and cardiac output. A pressure volume-loop catheter was placed into the left ventricle (LV) and provided continuous pressure and volume data of the LV, which allowed the calculation of both stroke volume and cardiac output.All three methods demonstrated good correlation with each other. The PV-loop catheter and thermodilution exhibited the best correlation with a 3% error and a Pearson coefficient of 0.99, with 95% CI=0.97 to 1.1, (p=0.002). The PV-loop catheter against ventriculography also showed good correlation with a 6% error and a Pearson coefficient of 0.95, 95% CI=0.96 to 1.1 (p=0.01). Finally, thermodilution against ventriculography had a 2% error with r=0.95, 95% CI=0.93 to 1.11, (p=0.01). In conclusion, we state that the PV-loop catheter, contrast ventriculography, and thermodilution each offer certain advantages depending on the researcher's requirements. Each method is reliable and accurate for measuring various cardiac parameters in swine such as the stroke volume and cardiac output.

Thermodilution, pressure-volume loop catheters, and contrast ventriculography are reliable and accurate methods for determining cardiac physiology such as stroke volume and cardiac output in a laboratory setting in swine.

Swine are frequently used in medical research given their similar cardiac physiology to that of humans. Measuring cardiac parameters such as stroke volume ...

Closed Chest Biventricular Pressure-Volume Loop Recordings with Admittance Catheters in a Porcine Model

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular evaluation is often performed in preclinical research. However, the right and left ventricles exert functional interdependence due to their parallel and serial connections, encouraging simultaneous evaluation of both ventricles. Furthermore, various pharmacological interventions may affect the ventricles and their preloads and afterloads differently. 
We describe our closed chest approach to admittance-based bi-ventricular PV loop recordings in a porcine model of acute right ventricular (RV) overload. We utilize minimally invasive techniques with all vascular accesses guided by ultrasound. PV catheters are positioned, under fluoroscopic guidance, to avoid thoracotomy in animals, as the closed chest approach maintains the relevant cardiopulmonary physiology. The admittance technology provides real-time PV loop recordings without the need for post-hoc processing. Furthermore, we explain some essential troubleshooting steps during critical timepoints of the presented procedure. 
The presented protocol is a reproducible and physiologically relevant approach to obtain a bi-ventricular cardiac PV loop recording in a large animal model. This can be applied to a large variety of cardiovascular animal research.

Here we present a closed chest approach to admittance-based bi-ventricular pressure-volume loop recordings in pigs with acute right ventricular dysfunction.

Pressure-volume (PV) loop recording enables the state-of-the-art investigation of load-independent variables of ventricular performance. Uni-ventricular ...

High-Resolution Three-Dimensional Imaging of the Footpad Vasculature in a Murine Hindlimb Gangrene Model

Peripheral arterial disease (PAD) is a significant cause of morbidity resulting from chronic exposure to atherosclerotic risk factors. Patients suffering from its most severe form, chronic limb-threatening ischemia (CLTI), face substantial impairments to daily living, including chronic pain, limited walking distance without pain, and nonhealing wounds. Preclinical models have been developed in various animals to study PAD, but mouse hindlimb ischemia remains the most widely used. There can be significant variation in response to ischemic insult in these models depending on the mouse strain used and the site, number, and means of arterial disruption. This protocol describes a unique method combining femoral artery and vein electrocoagulation with the administration of a nitric oxide synthase (NOS) inhibitor to reliably induce footpad gangrene in Friend Virus B (FVB) mice that resembles the tissue loss of CLTI. While traditional means of assessing reperfusion such as laser Doppler perfusion imaging (LDPI) are still recommended, intracardiac perfusion of the lipophilic dye 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI) is used to label the vasculature. Subsequent whole-mount confocal laser scanning microscopy allows for high-resolution, three-dimensional (3D) reconstruction of footpad vascular networks that complements traditional means of assessing reperfusion in hindlimb ischemia models.

The present protocol describes a unique, clinically relevant model of peripheral arterial disease that combines femoral artery and vein electrocoagulation with the administration of a nitric oxide synthase inhibitor to induce hindlimb gangrene in FVB mice. Intracardiac DiI perfusion is then used for high-resolution, three-dimensional imaging of the footpad vasculature.

Peripheral arterial disease (PAD) is a significant cause of morbidity resulting from chronic exposure to atherosclerotic risk factors. Patients suffering ...

Four-Dimensional Computed Tomography-Guided Valve Sizing for Transcatheter Pulmonary Valve Replacement

The measurements of the right ventricle (RV) and pulmonary artery (PA), for selecting the optimal prosthesis size for transcatheter pulmonary valve replacement (TPVR), vary considerably. Three-dimensional (3D) computed tomography (CT) imaging for device size prediction is insufficient to assess the displacement of the right ventricular outflow tract (RVOT) and PA, which could increase the risk of stent misplacement and paravalvular leak. The aim of this study is to provide a dynamic model to visualize and quantify the anatomy of the RVOT to PA over the entire cardiac cycle by four-dimensional (4D) cardiac CT reconstruction to obtain an accurate quantitative evaluation of the required valve size. In this pilot study, cardiac CT from sheep J was chosen to illustrate the procedures. 3D cardiac CT was imported into 3D reconstruction software to build a 4D sequence which was divided into eleven frames over the cardiac cycle to visualize the deformation of the heart. Diameter, cross-sectional area, and circumference of five imaging planes at the main PA, sinotubular junction, sinus, basal plane of the pulmonary valve (BPV), and RVOT were measured at each frame in 4D straightened models prior to valve implantation to predict the valve size. Meanwhile, dynamic changes in the RV volume were also measured to evaluate right ventricular ejection fraction (RVEF). 3D measurements at the end of the diastole were obtained for comparison with the 4D measurements. In sheep J, 4D CT measurements from the straightened model resulted in the same choice of valve size for TPVR (30 mm) as 3D measurements. The RVEF of sheep J from pre-CT was 62.1 %. In contrast with 3D CT, the straightened 4D reconstruction model not only enabled accurate prediction for valve size selection for TPVR but also provided an ideal virtual reality, thus presenting a promising method for TPVR and the innovation of TPVR devices.

This study assessed a new methodology with a straightened model generated from the four-dimensional cardiac computed tomography sequence to obtain the desired measurements for valve sizing in the application of transcatheter pulmonary valve replacement.

The measurements of the right ventricle (RV) and pulmonary artery (PA), for selecting the optimal prosthesis size for transcatheter pulmonary valve ...

Drug Treatment by Central Venous Catheter in a Mouse Model of Angiotensin II Induced Abdominal Aortic Aneurysm and Monitoring by 3D Ultrasound

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse models, are applied to advance the understanding of the disease pathogenesis and to identify potential therapeutic targets. Testing novel drug candidates to block AAA growth in these models generally requires repeated drug administration during the time course of the experiment. Here, we describe a compiled protocol for AAA induction, insertion of an intravenous catheter to facilitate prolonged therapy, and serial AAA monitoring by 3D ultrasound. Aneurysms are induced in apolipoprotein E (ApoE) deficient mice by angiotensin II release over 28 days from osmotic mini-pumps implanted subcutaneously into the mouse back. Subsequently, the surgical procedure for external jugular vein catheterization is conducted to allow for daily intravenous drug treatment or repeated blood sampling via a subcutaneous vascular access button. Despite the two dorsal implants, the monitoring of AAA development is readily facilitated by sequential semi-automated 3D ultrasound analysis, which yields comprehensive information on the expansion of aortic diameter and volume and on aneurysm morphology, as illustrated by experimental examples.

This protocol describes the consecutive implantation of an osmotic pump to induce abdominal aortic aneurysm by angiotensin II release in apolipoprotein E (ApoE) deficient mice and of a vascular access port with a jugular vein catheter for repeated drug treatment. Monitoring of aneurysm development by 3D ultrasound is effectively conducted despite dorsal implants.

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse ...

Image Acquisition Method for the Sonographic Assessment of the Inferior Vena Cava

Over the past several decades, clinicians have incorporated several applications of diagnostic point-of-care ultrasound (POCUS) into medical decision-making. Among the applications of POCUS, imaging the inferior vena cava (IVC) is practiced by a wide variety of specialties, such as nephrology, emergency medicine, internal medicine, critical care, anesthesiology, pulmonology, and cardiology. Although each specialty uses IVC data in slightly different ways, most medical specialties, at minimum, attempt to use IVC data to make predictions about intravascular volume status. While the relationship between IVC sonographic data and intravascular volume status is complex and highly context-dependent, all clinicians should collect the sonographic data in standardized ways to ensure repeatability. This paper describes standardized IVC image acquisition including patient positioning, transducer selection, probe placement, image optimization, and the pitfalls and limitations of IVC sonographic imaging.&#160;This paper also describes the commonly performed anterior IVC long-axis view and three other views of the IVC that can each provide helpful diagnostic information when the anterior long-axis view is difficult to obtain or interpret.

Point-of-care ultrasound evaluation of the inferior vena cava (IVC) is commonly utilized to identify, among other things, the volume status. Imaging should be performed systematically to ensure repeatability. This manuscript reviews the methods and pitfalls of sonographic IVC examination.

Over the past several decades, clinicians have incorporated several applications of diagnostic point-of-care ultrasound (POCUS) into medical ...

Modified Arteriovenous Fistula Surgery to Establish the RV Volume Overload Mouse Model

Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model

Right ventricular (RV) volume overload (VO) is common in children with congenital heart disease. In view of distinct developmental stages,the RV myocardium may respond differently to VO in children compared to adults. The present study aims to establish a postnatal RV VO model in mice using a modified abdominal arteriovenous fistula. To confirm the creation of VO and the following morphological and hemodynamic changes of the RV, abdominal ultrasound, echocardiography, and histochemical staining were performed for 3 months. As a result, the procedure in postnatal mice showed an acceptable survival and fistula success rate. In VO mice, the RV cavity was enlarged with a thickened free wall, and the stroke volume was increased by about 30%-40% within 2 months after surgery. Thereafter, the RV systolic pressure increased, corresponding pulmonary valve regurgitation was observed, and small pulmonary artery remodeling appeared. In conclusion, modified arteriovenous fistula (AVF) surgery is feasible to establish the RV VO model in postnatal mice. Considering the probability of fistula closure and elevated pulmonary artery resistance, abdominal ultrasound and echocardiography must be performed to confirm the model status before application.

Author Spotlight: Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model  

This protocol presents the establishment and confirmation of a postnatal right ventricular volume overload (VO) model in mice with abdominal arteriovenous fistula (AVF), which can be applied to investigate how VO contributes to postnatal heart development.

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

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