Name: femoral arterial and venous catheterization for blood sampling, drug administration and conscious blood pressure and heart rate measurements
Uploaded: 2012-01-24T12:00:00
Duration: 9 min 38 s
Description: Watch this Scientific Journal Video about Femoral Arterial and Venous Catheterization for Blood Sampling, Drug Administration and Conscious Blood Pressure and Heart Rate Measurements at JoVE.com

The authors wish to acknowledge Dr. Andrew King, PhD and Katrina King for the contribution of the illustrations. NIH grant: R00HL087927.

1. Prior to Starting Surgical Procedure
Note: Prior to beginning any animal procedures ensure that you have obtained the appropriate permission through your institution/organization. As with all survival surgeries, make sure that the surgery is performed under sterile conditions and the appropriate pain medication and antibiotics needed are used for a successful outcome.
<ol>
	<li>Prior to beginning the catheterization procedure evaluate the catheters by flushing them with sterile saline to ensure they are patent.</li>
	<li>Anesthetize the rat.</li>
	<li>Prepare the rat for the surgical procedure.
	<ol class="lalpha">
		<li>Shave the fur from the surgical regions, which include the back of the neck (between the shoulder blades) and the inner leg region.</li>
		<li>Using Betadine and a 70% ethanol scrub respectively, scrub the shaved surgical regions starting in the center and making a circular sweep outwards. Repeat this 3 times for each region, finishing with a final cleansing with 70% ethanol.</li>
		<li>Place the animal on a sterile surface and place sterile drape over surgical areas. (The use of Press-n-Seal is an ideal surgical draping allowing the investigator to monitor the animal throughout the procedure.)</li>
	</ol>
	</li>
	<li>Ensure all surgical tools for the surgical procedure have been sterilized.</li>
</ol>
2. Preparation of Surgical Areas for Catheter Placement
<ol>
	<li>With the rat laying prone (on its stomach), make approximately a &frac12; inch (12 mm) horizontal incision on the back of the neck at the level of the shoulder blades with scissors or a scalpel, then blunt dissect a subcutaneous "pocket" in the back approximately the size of a quarter. This will be used as an area to place a small amount of tubing that will compensate for animal growth and/or movement, i.e. so that the catheter isn't pulled on and thus removed from the artery it was placed in. (Alternatively this step can be performed immediately prior to tunneling the catheter.)</li>
	<li>Place the rat onto its back (supine position) and make an incision in the inguinal area [approximate &frac12; inch (12 mm) incision along the natural angle of the hind leg.</li>
	<li>Blunt dissect to separate the connective tissue (Figure 1) (can use blunt-tip scissors, hemostats, cotton swabs, etc.) (typically by holding your blunt tipped scissors and/or cotton swabs at a 45 degree angle this ensures the easier localization of the region of interest) until the femoral artery and vein are exposed.
	<ol class="lalpha">
		<li>The vein is dark red in color and the artery is clearer and brighter than the vein. The nerve that runs along the artery is whitish in tone.</li>
	</ol>
	</li>
	<li>Blunt dissect in the leg region to make a small open region below the skin (i.e. pocket, approximately the size of a quarter) along the inside of the leg for placement of a small section of the catheter (again to take into account animal movement and growth of the animal if chronic placement).</li>
	<li>Place the retractors into the incision area so that you can fully view the artery and vein.</li>
	<li>Using fine tip forceps gently separate the nerve (whitish in color) that is along the femoral artery away from the artery and vein. (Figure 2) Be careful to not cut or damage the nerve.</li>
	<li>Separate the artery and vein as one unit, trying to expose an approximately &frac14; inch (5-7 mm) length section of artery/vein.</li>
	<li>Repeat the process in separating the vein from the artery. Keep your surgical instruments (i.e. fine tip forceps) perpendicular to the vessels and separate the vessels in parallel. This assists in avoiding tearing, pricking or damaging the vessels. By placing the fine tip forceps gently between the artery and vein from underneath and slowly opening the forceps and repeating this, you will slowly separate the vessels. Note: if you do tear or observe some bleeding use a sterile cotton swab and/or 2 x 2 gauze and place pressure on the area until the bleeding has stopped, then continue with the surgery.</li>
</ol>
3. Tunneling the Catheter
<ol>
	<li>Place sterile saline soaked 2 x 2 gauze over the incision and turn the animal to its stomach.</li>
	<li>Place the Rochester Pean forceps (long straight forceps) into the incision on the back that was made previously and guide the forceps subcutaneously down the back to the level of the hips [ensure the tips of the hemostats are pointed up, (not towards the spine) to avoid injuring the spinal cord]. At approximately the hip region turn the hemostat tip towards the incision that was made in the leg region and push the tip of the hemostats out of the prepared leg incision.</li>
	<li>Gently grasp the end of the catheters (not the end that will be inserted into the artery/vein) with the forceps and gently pull the catheters through the cavity that was made and ultimately out the neck incision.</li>
	<li>Place the appropriate blunted-tip needle syringe filled with 20 U/ml Heparin/saline on the end of the respective catheters and fill the catheters (ensure that there are no air bubbles) with the heparinized/saline. Once again check that the catheters are patent and no damage has come to the catheters. Leave the syringes attached to the end of the catheter lines to ensure no air is able to get into the catheter lines and turn the rat onto its back once again.</li>
</ol>
4. Inserting the Catheters into the Femoral Vein and Artery (Figure 3)
<ol>
	<li>Place a folded piece of sterile 4.0 silk/thread under the femoral vein and then cut the silk at the folded end. There will now be two pieces of silk under the vein. Or alternatively, place 2 pieces of 4.0 silk under the vein individually.</li>
	<li>Separate the silk that you placed under the vein, one piece towards the leg (distal end) and the other towards the body. Tie a loose ligature on the side closest to the body, then using small hemostats, grasp the silk and pull taught but do not tighten the knot. Pull the other piece of silk as far as possible towards the leg (distal end) and tie this into a triple knot, grasp the silk with small hemostats and pull taught. This method will allow the vein to fill with blood, making it easier to make the the incision that is required for inserting the catheter (step 4).</li>
	<li>Place 1-2 drops of lidocaine onto the vein.</li>
	<li>Using the Vanna micro-dissecting scissors, make a small incision in the vein approximately &frac14; thru and at a 45 degree angle.</li>
	<li>Place fine-tipped forceps (45 forceps) into the incision and using another pair of forceps; carefully feed in the vein catheter. Gently open the forceps that are placed into the vein, as this will allow the researcher to gently place the venous catheter under the forceps and into the vein.</li>
	<li>When the catheter is fully inserted (approx. 6-7 cm) (when making the catheter a mark is placed on the catheter to assist the surgeon in identifying when the catheter is fully inserted) (this places the venous catheter in the abdominal vena cava), tighten the anterior ligature around the vein and catheter, tying a triple knot (ensure that it isn't occluding the vein). Use the silk suture near the leg (posterior ligature) to again secure the catheter (triple knot) and ensure placement. Slowly draw back the syringe until there is a little blood visible in the catheter, which helps to ensure that the suture knots are not too tight and that the catheter is functional. After checking, depress the plunger until the blood is no longer visible in the catheter. 
	Note: it is possible to insert 2 catheters into the femoral vein if needed.</li>
	<li>Repeat steps 9-13 to place a femoral artery catheter with the following exceptions:
	<ol class="lalpha">
		<li>Tie the silk nearest to the leg (posterior) with a triple knot and pull taught before tying the loose ligature near the body (anterior) prior to making the incision for the catheter placement. This will allow the artery to fill with blood making it easier to cut. Ensure the proximal suture is pulled taught to occlude the artery prior to cutting the artery. This will avoid blood loss when the cut is made.</li>
		<li>Insert the arterial catheter approximately 5 cm from the femoral artery (this places the catheter in the abdominal aorta).</li>
		<li>When securing the catheter with the suture, make sure the suture is not too tight and occluding the catheter.</li>
	</ol>
	</li>
</ol>
5. Surgical Wrap-up
<ol>
	<li>Make a dime to quarter sized loop in the catheters and place on the inside of the leg (the loop should fit in the area that was blunt dissected earlier). After placing both catheters, secure them with 1-2 stitches of 5.0 surgical suture into the muscle layer.</li>
	<li>Close the incision with 4.0 Ethilon with non-continuous sutures.</li>
	<li>Turn the rat onto its stomach and make another loop in the catheters about the size of a quarter and place in the pocket dissected in the back. Close the incision with suture.</li>
	<li>Using a drop of vetbond, secure the catheters in the back.</li>
	<li>Clamp the catheters near the back incision with padded hemostats and remove the syringes from the ends.</li>
	<li>Fit the rat with a tether-type jacket, cap the catheters (to maintain the heparin lock), and remove the padded hemostats.</li>
</ol>
--For long-term maintenance replace catheter saline solution with 20 U/ml heparin/saline.
6. Maintenance of Catheter (Sterile gloves should be worn during procedure)
<ol>
	<li>Clamp the catheter with padded forceps.</li>
	<li>Remove the catheter plug.</li>
	<li>Place a blunt tipped syringe with lock solution onto the catheter.</li>
	<li>Unclamp the forceps.</li>
	<li>Fill the catheter with the lock solution (volume is pre-determined-typically 0.3 ml).</li>
	<li>Clamp the catheter while flushing to prevent any backflow of blood into the catheter tip and remove the syringe.</li>
	<li>Replace the catheter plug.</li>
	<li>Unclamp the forceps and gently push the plug in slightly to ensure no blood is in the tip of the catheter.</li>
</ol>
7. Blood Sampling (Sterile gloves should be worn during procedure)
Clamp the catheter with padded forceps.
<ol>
	<li>Remove the catheter plug.</li>
	<li>Slowly withdraw lock solution using a blunt tipped syringe and discard.</li>
	<li>Attach sampling syringe to the catheter and slowly withdraw the sample.</li>
	<li>Clamp the catheter with padded forceps and place a syringe with lock solution onto the catheter and re-fill the catheter with the lock solution.</li>
	<li>Clamp the catheter while re-filling the catheter with the solution.</li>
	<li>Re-insert the catheter plug.</li>
	<li>Remove the padded forceps and gently push the plug in slightly further.</li>
</ol>
8. Drug Infusion
<ol>
	<li>Clamp the catheter with padded forceps.</li>
	<li>Remove the catheter plug.</li>
	<li>Slowly withdraw lock solution using a blunt tipped syringe and discard.</li>
	<li>Attach drug filled syringe to the catheter and infuse the substance into the animal.
	<ul>
		<li>one can use a 3 way stopcock as well as an intermediate if multiple injections are required, thus requiring less fluid infusion into the animal.</li>
		<li>one can also attach a constant infusion syringe pump with a sterile filter for continuous infusions.</li>
	</ul>
	</li>
	<li>Clamp with padded forceps and place a syringe with lock solution onto the catheter and re-fill the catheter with the lock solution.</li>
	<li>Clamp the catheter while re-filling the catheter with the solution.</li>
	<li>Re-insert the catheter plug.</li>
	<li>Remove the padded forceps and gently push the plug in slightly further.</li>
</ol>
9. Blood Pressure and Heart Rate Sampling
<ol>
	<li>Clamp the arterial catheter with padded forceps and remove the catheter plug.</li>
	<li>Attach the arterial line to the pressure transducer.</li>
	<li>Follow manufacture instructions for using the software for the blood pressure collection.</li>
	<li>At the conclusion of the blood pressure monitoring period, using padded forceps clamp the catheter and disconnect from the transducer.</li>
	<li>Flush the catheter with the lock solution and replace the catheter plug as described above.</li>
</ol>
10. Representative Results
A representative blood pressure measure was taken from a conscious freely moving animal and is presented in Figure 4. Phenylephrine (3 ug/kg, iv), an alpha 1 adrenergic receptor agonist, was administered into the femoral vein line to increase blood pressure, while simultaneously measuring blood pressure from the femoral arterial line. Phentolamine (4 mg/kg, iv), a nonselective alpha-adrenergic antagonist, was then administered to lower blood pressure.
<img alt="Figure 1" src="/files/ftp_upload/3496/3496fig1.jpg" /> 
Figure 1. Blunt dissection of tissue. With the rat on his back, you will have made approximately a &frac12; inch (12 mm) incision on the angle of the hind leg and using blunt scissors or hemostats you will blunt dissect the connective tissue to expose the femoral artery and vein.
<img alt="Figure 2" src="/files/ftp_upload/3496/3496fig2.jpg" /> 
Figure 2. Separation of femoral artery and vein from connective tissue.
<img alt="Figure 3" src="/files/ftp_upload/3496/3496fig3.jpg" /> 
Figure 3. Catheter placement. Using Vanna micro-dissecting scissors place a small incision approximately &frac14; of the way through the vessel at a 45 degree angle (top) and place fine-tipped forceps into the incision and using another pair of forceps feed the catheter into the vessel (middle). Lastly, upon completion of the placement of the catheter suture the catheter in place (bottom).
<img alt="Figure 4" src="/files/ftp_upload/3496/3496fig4.jpg" /> 
Figure 4. Representative blood pressure measure taken from a conscious freely moving animal.

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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+arterial+and+venous+access+in+the+unrestrained+rat.">Chronic arterial and venous access in the unrestrained rat.</a> Am. J. Physiol. 238, H599-H603 (1980).</li><li>Cocchetto, D. M., Bjornsson, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+vascular+access+and+collection+of+body+fluids+from+the+laboratory+rat.">Methods for vascular access and collection of body fluids from the laboratory rat.</a> J. Pharm. Sci. 72, 465-492 (1983).</li><li>Rigalli, A., E, D. L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Experimental surgical models in the laboratory rat. , (2009).</li><li>Waynforth, H. 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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+analysis+of+ACTH+and+corticosterone+sampling+methods+in+rats.">Comparative analysis of ACTH and corticosterone sampling methods in rats.</a> Am. J. Physiol. Endocrinol. Metab. 289, E823-E828 (2005).</li><li>Buhler, H. U., da Prada, M., Haefely, W., Picotti, G. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma+adrenaline,+noradrenaline+and+dopamine+in+man+and+different+animal+species.">Plasma adrenaline, noradrenaline and dopamine in man and different animal species.</a> J. Physiol. 276, 311-320 (1978).</li><li>Fagin, K. D., Shinsako, J., Dallman, M. 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J., Mann, M., Osborn, J., Werber, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continuous+blood+pressure+measurement+in+rats+with+aortic+baroreceptor+deafferentation.">Continuous blood pressure measurement in rats with aortic baroreceptor deafferentation.</a> Am. J. Physiol. 241, H268-H272 (1981).</li><li>Van Vliet, B. N., Chafe, L. L., Antic, V., Schnyder-Candrian, S., Montani, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+and+indirect+methods+used+to+study+arterial+blood+pressure.">Direct and indirect methods used to study arterial blood pressure.</a> J. Pharmacol. Toxicol Methods. 44, 361-373 (2000).</li><li>Wang, J., Tempini, A., Schnyder, B., Montani, J. 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Production and Free Access of this video-article is sponsored by ADInstruments.

Arterial and vein catheterization have historically been used to both acutely and chronically monitor blood pressure, sample blood and deliver substances in the experimental rat animal model1-4. A major benefit of these surgical instrumentations is that it allows monitoring procedures, including, blood sampling, drug administration and blood pressure monitoring, to be conducted with minimal disturbances and/or stress to the animal1. Numerous investigators have written protocols and have specific methodologies by which their laboratory performs the surgical procedure5-8. The video and illustrations demonstrate what our laboratory has found to be successful with regards to the femoral arterial and venous catheterization procedure.
Rats are commonly used in the laboratory for a multitude of scientific studies due to their small size and convenience in handling. There are several locations where a chronic catheter can be placed within an animal, including the jugular vein, abdominal aorta, carotid artery and femoral artery, to name a few. The rat femoral location for chronic catheterization results in increased length of catheter patency and had the fastest recovery of pre-surgical animal weight compared to other catheter locations9.
Numerous investigators have used indwelling catheters for acute as well as chronic blood sampling1-5,11-13. In many studies multiple blood draws are required of an animal and external cannulation/catheterization is one methodology that is advantageous due to its non-traumatic nature; moreover it can be done while the animal is conscious, thus is not limited by the effects of anesthetics and also the animal can be freely moving1,10. The best method by which to obtain blood samples and to measure stress hormones in rodents has been long debated6,13. With regards to pharmacokinetic studies, catheterization of the research animal permits repeated blood sampling with minimal restraint when using a chronically implanted catheter system. In addition, studies have shown reduced basal concentrations of adrenaline, noradrenaline and dopamine in the plasma of freely moving rats (jugular catheter) compared to animals that have been handled (30 seconds) and or restrained (5 minutes)14. An additional method by which to discern increases in stress is by measuring plasma corticosterone levels. Previously it was suggested that even following three to four days of surgical recovery times that plasma corticosterone levels were elevated in the chronic cannulated rat15. However, recent improvements in methodology have determined that there are no differences in baseline plasma corticosterone levels in the jugular vein cannulated rats compared to uncannulated rats16. In addition, HPLC methodology for corticosterone analysis also revealed that corticosterone levels are elevated by restraint stress; however, stable after jugular catheterization16.
An additional use of chronic catheterization is the measurement of blood pressure and heart rate in rodents. There are multiple methodologies that are utilized to measure the blood pressure and heart rate in the rat; these include the non-invasive tail cuff methodologies, radiotelemetry procedures and direct indwelling catheters. Each method has its advantages and disadvantages, which are described in detail in other publications. Indwelling fluid-filled catheters can be implanted into multiple arteries within the rat. The femoral artery is but one artery that can be used for this measure. For blood pressure/heart rate measures, the distal end of the catheter is connected to a calibrated pressure transducer . The catheter can be housed in a protective spring that is connected to a swivel to allow free movement of the animal, or attached to a button surgically implanted to the animal. Indwelling catheters have the advantage of minimizing the long-term stress on the animal17,18. In addition the materials are inexpensive, calibration is easy for pressure measures and continuous long-term measures can be obtained under conditions of relatively low stress for numerous weeks19. We would be remiss if we didn't mention that there are disadvantages to this technique, including, it is a surgically invasive technique, maintenance of the catheter is required to maintain patency, damage to the artery due to the implantation of the catheter and the potential of infection to name a few.
For chronic measures of blood pressure direct recordings from chronically-implanted arterial catheters are more technically challenging but are more accurate and can be done continuously without disturbing the animal. Tail-cuff measurements are less accurate; however they do not require surgery and can also be repeated. Tail-cuff methodologies do require handling as well as heating of the animal to dilate the tail vessels to facilitate the detection of the pulse20. Handling and the added heat stress can affect blood pressure measurements, thus not providing truly accurate measures. Moreover, the non-direct tail-cuff method does not permit easy simultaneous blood sampling or drug administration.
An additional method by which direct measure can be achieved is using telemetric methodology. Telemetry allows high-quality recordings of blood pressure (and other measures) continuously for long periods of time in conscious freely moving animals without restraint or anesthetics18. However, telemetry devices while ideal are very costly. When compared to telemetry, catheterization benefits include: decreased "setup" and operational costs, the ability to readily administer drugs and easily take blood samples in conscious freely moving animals. The administration of drugs and substances, and obtaining blood samples from the research animal can be done while minimally disturbing the animal, thus minimizing stress to the animal and allowing for a more accurate measure.

<table border="1">
 <tbody>
 <tr>
 <td>Name</td>
 <td>Type</td>
 <td>Company</td>
 <td>Catalog Number</td>
 </tr>
 <tr>
 <td>Arterial Catheter</td>
 <td>Consumable</td>
 <td>SAI Infusion</td>
 <td>RFA-01</td>
 </tr>
 <tr>
 <td>Venous Catheter</td>
 <td>Consumable</td>
 <td>Scientific Commodities, Inc. </td>
 <td>BB518-20 &amp; BB31785-V/2</td>
 </tr>
 <tr>
 <td>27g 1/2" Blunt Needle</td>
 <td>Equipment</td>
 <td>Integrated Dispensing Solutions, Inc.</td>
 <td>9991113</td>
 </tr>
 <tr>
 <td>23g 1/2" Blunt Needle</td>
 <td>Equipment</td>
 <td>Integrated Dispensing Solutions, Inc.</td>
 <td>9991109</td>
 </tr>
 <tr>
 <td><a href="http://www.adinstruments.com/products/hardware/research/product/PL3508/" target="_blank">PowerLab/8SP</a></td>
 <td>Equipment</td>
 <td><a href="http://www.adinstruments.com/" target="_blank">ADInstruments</a></td>
 <td><a href="http://www.adinstruments.com/products/hardware/research/product/PL3508/" target="_blank">ML765</a></td>
 </tr>
 <tr>
 <td><a href="http://www.adinstruments.com/products/hardware/research/product/FE224/" target="_blank">Quad Bridge</a></td>
 <td>Equipment</td>
 <td><a href="http://www.adinstruments.com/" target="_blank">ADInstruments</a></td>
 <td><a href="http://www.adinstruments.com/products/hardware/research/product/FE224/" target="_blank">ML118</a></td>
 </tr>
 <tr>
 <td><a href="http://www.adinstruments.com/products/software/research/LabChart-Software/" target="_blank">LabChart 7.2</a></td>
 <td>Software</td>
 <td><a href="http://www.adinstruments.com/" target="_blank">ADInstruments</a></td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td><a href="http://www.adinstruments.com/products/hardware/research/product/MLT844/" target="_blank">BP Transducer with stopcock &amp; Cable</a></td>
 <td>Equipment</td>
 <td><a href="http://www.adinstruments.com/" target="_blank">ADInstruments</a></td>
 <td><a href="http://www.adinstruments.com/products/hardware/research/product/MLT844/" target="_blank">MLT844</a></td>
 </tr>
 <tr>
 <td>Single Channel Swivel</td>
 <td>Equipment</td>
 <td>Instech</td>
 <td>375/22PS</td>
 </tr>
 <tr>
 <td>Single Axis-Counter Balance Swivel Mount</td>
 <td>Equipment</td>
 <td>Instech</td>
 <td>CM375BP</td>
 </tr>
 <tr>
 <td>Covance Infusion Harness</td>
 <td>Equipment</td>
 <td>Instech</td>
 <td>CIH105</td>
 </tr>
 <tr>
 <td>Table Top Isoflurane Anesthesia Unit</td>
 <td>Equipment</td>
 <td>Protech International, Inc.</td>
 <td>61020</td>
 </tr>
 <tr>
 <td>Rodent Anesthesia Mask</td>
 <td>Equipment</td>
 <td>Protech International, Inc.</td>
 <td>RAM-02</td>
 </tr>
 <tr>
 <td>Rodent Induction Chamber</td>
 <td>Equipment</td>
 <td>Protech International, Inc.</td>
 <td>RIC-01</td>
 </tr>
 <tr>
 <td>Anesthesia Gas Filter Canister</td>
 <td>Equipment</td>
 <td>Protech International, Inc.</td>
 <td>80120</td>
 </tr>
 <tr>
 <td>Tabletop Laminar Flow Hood</td>
 <td>Equipment</td>
 <td>Sentry Air Systems, Inc.</td>
 <td>SS-200-WSL</td>
 </tr>
 <tr>
 <td>Oster Golden A5</td>
 <td>Equipment</td>
 <td>Oster Professional Products</td>
 <td>78005-140</td>
 </tr>
 <tr>
 <td>Fiber Optic Light with Dual Gooseneck Guide</td>
 <td>Equipment</td>
 <td>LW Scientific, Inc. </td>
 <td>ILL-1502-DGG1</td>
 </tr>
 <tr>
 <td>Deltaphase Isothermal Pad</td>
 <td>Equipment</td>
 <td>BrainTree Scientific</td>
 <td>39DP</td>
 </tr>
 <tr>
 <td>Cotton-Tipped Applicators</td>
 <td>Consumable</td>
 <td>Solon Manufacturing Company</td>
 <td>36200</td>
 </tr>
 <tr>
 <td>Gauze Sponges 2"x2"</td>
 <td>Consumable</td>
 <td>Kendall Curity</td>
 <td>2146</td>
 </tr>
 <tr>
 <td>5-0 Nylon Suture</td>
 <td>Consumable</td>
 <td>Ethicon</td>
 <td>661G</td>
 </tr>
 <tr>
 <td>4-0 Silk Suture</td>
 <td>Consumable</td>
 <td>Deknatel</td>
 <td>8-S, 136075-0208</td>
 </tr>
 <tr>
 <td>Tissue Adhesive</td>
 <td>Consumable</td>
 <td>3M</td>
 <td>1469SB</td>
 </tr>
 <tr>
 <td>Splinter &amp; Fixation Forceps</td>
 <td>Equipment</td>
 <td>George Tiemann &amp; Company</td>
 <td>160-55</td>
 </tr>
 <tr>
 <td>Student Tissue Forceps</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>91121-12</td>
 </tr>
 <tr>
 <td>Micro Forceps/Bracken Forceps</td>
 <td>Equipment</td>
 <td>George Tiemann &amp; Company</td>
 <td>10-1942</td>
 </tr>
 <tr>
 <td>Extra Fine Graefe Forceps</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>11152-10</td>
 </tr>
 <tr>
 <td>Olsen-Hegar Needle Holder</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>12002-12</td>
 </tr>
 <tr>
 <td>Student Halsted-Mosquito Hemostat</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>91308-12</td>
 </tr>
 <tr>
 <td>Rochester Pean Forcep </td>
 <td>Equipment</td>
 <td>Biomedical Research Instruments, Inc.</td>
 <td>31-1640</td>
 </tr>
 <tr>
 <td>Student Surgical Scissors</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>91402-12</td>
 </tr>
 <tr>
 <td>Dumont Forceps</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>11251-35</td>
 </tr>
 <tr>
 <td>Micro Dissecting Scissors</td>
 <td>Equipment</td>
 <td>George Tiemann &amp; Company</td>
 <td>160-210</td>
 </tr>
 <tr>
 <td>Strabismus Scissors</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>14075-11</td>
 </tr>
 <tr>
 <td>Hartman Hemostat</td>
 <td>Equipment</td>
 <td>Fine Science Tools, Inc.</td>
 <td>13003-10</td>
 </tr>
 <tr>
 <td>Tissue Scissors</td>
 <td>Equipment</td>
 <td>George Tiemann &amp; Company</td>
 <td>160-150</td>
 </tr>
 <tr>
 <td>Retractor</td>
 <td>Equipment</td>
 <td>Custom Made</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

femoral arterial and venous catheterization for blood sampling, drug administration and conscious blood pressure and heart rate measurements

In multiple fields of study, access to the circulatory system in laboratory studies is necessary. Pharmacological studies in rats using chronically implanted catheters permit a researcher to effectively and humanely administer substances, perform repeated blood sampling and assists in conscious direct measurements of blood pressure and heart rate. Once the catheter is implanted long-term sampling is possible. Patency and catheter life depends on multiple factors including the lock solution used, flushing regimen and catheter material. This video will demonstrate the methodology of femoral artery and venous catheterization of the rat. In addition the video will demonstrate the use of the femoral venous and arterial catheters for blood sampling, drug administration and use of the arterial catheter in taking measurements of blood pressure and heart rate in a conscious freely-moving rat. A tether and harness attached to a swivel system will allow the animal to be housed and have samples taken by the researcher with minimal disruption to the animal. To maintain patency of the catheter, careful daily maintenance of the catheter is required using lock solution (100 U/ml heparinized saline), machine-ground blunt tip syringe needles and the use of syringe filters to minimize potential contamination. With careful aseptic surgical techniques, proper catheter materials and careful catheter maintenance techniques, it is possible to sustain patent catheters and healthy animals for long periods of time (several weeks).

Watch this Scientific Journal Video about Femoral Arterial and Venous Catheterization for Blood Sampling, Drug Administration and Conscious Blood Pressure and Heart Rate Measurements at JoVE.com

Femoral Arterial and Venous Catheterization for Blood Sampling, Drug Administration and Conscious Blood Pressure and Heart Rate Measurements

Chronic catheterization of blood vessels in the rat is often required for administration of substances, obtain blood sample over a period of time or for direct conscious blood pressure measurements. Femoral arterial catheterization of the rat and corresponding measurements of blood pressure in the conscious animal will be demonstrated.

In multiple fields of study, access to the circulatory system in laboratory studies is necessary. Pharmacological studies in rats using chronically ...

femoral-arterial-venous-catheterization-for-blood-sampling-drug

Research

JoVE Journal

Medicine

Catheterization of the Carotid Artery and Jugular Vein to Perform Hemodynamic Measures, Infusions and Blood Sampling in a Conscious Rat Model

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. Intra-tracheal inoculation of a known amount of bacteria has been successfully used to reproduce the pathogenesis of pneumonia which then develops into sepsis. Monitoring hemodynamic parameters and providing standard clinical treatment including infusion of antibiotics, fluids and drugs to maintain blood pressure is critical to simulate routine supportive care in this model but to do so requires both arterial and venous vascular access. The video details the surgical technique for implanting carotid artery and common jugular vein catheters in an anesthetized rat. Following a 72 hr recovery period, the animals will be re-anesthetized and connected to a tether and swivel setup attached to the rodent housing which connects the implanted catheters to the hemodynamic monitoring system. This setup allows free movement of the rat during the study while continuously monitoring pressures, infusing fluids and drugs (antibiotics, vasopressors) and performing blood sampling.

Vascular accesses to measure hemodynamics, provide fluids and perform blood sampling are important to any small animal model study. We present a technique for implanting catheters into the carotid artery and the common jugular vein in an anesthetized rat for connecting to a system to perform monitoring, infusions and sampling.

The success of a small animal model to study critical illness is, in part, dependent on the ability of the model to simulate the human condition. ...

Quantification of the Immunosuppressant Tacrolimus on Dried Blood Spots Using LC-MS/MS

The calcineurin inhibitor tacrolimus is the cornerstone of most immunosuppressive treatment protocols after solid organ transplantation in the United States. Tacrolimus is a narrow therapeutic index drug and as such requires therapeutic drug monitoring and dose adjustment based on its whole blood trough concentrations. To facilitate home therapeutic drug and adherence monitoring, the collection of dried blood spots is an attractive concept. After a finger stick, the patient collects a blood drop on filter paper at home. After the blood is dried, it is mailed to the analytical laboratory where tacrolimus is quantified using high-performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) in combination with a simple manual protein precipitation step and online column extraction.
For tacrolimus analysis, a 6-mm disc is punched from the saturated center of the blood spot. The blood spot is homogenized using a bullet blender and then proteins are precipitated with methanol/0.2 M ZnSO4 containing the internal standard D2,13C-tacrolimus. After vortexing and centrifugation, 100 &#181;l of supernatant is injected into an online extraction column and washed with 5 ml/min of 0.1 formic acid/acetonitrile (7:3, v:v) for 1 min. Hereafter, the switching valve is activated and the analytes are back-flushed onto the analytical column (and separated using a 0.1% formic acid/acetonitrile gradient). Tacrolimus is quantified in the positive multi reaction mode (MRM) using a tandem mass spectrometer.
The assay is linear from 1 to 50 ng/ml. Inter-assay variability (3.6%-6.1%) and accuracy (91.7%-101.6%) as assessed over 20 days meet acceptance criteria. Average extraction recovery is 95.5%. There are no relevant carry-over, matrix interferences and matrix effects. Tacrolimus is stable in dried blood spots at RT and at +4 &#176;C for 1 week. Extracted samples in the autosampler are stable at +4 &#176;C for at least 72 hr.

Here we describe a high-performance liquid chromatography-tandem mass spectrometry (HPLC-MS/MS) assay to quantify the immunosuppressant tacrolimus in dried blood spots using a simple manual protein precipitation step and online column extraction.

The calcineurin inhibitor tacrolimus is the cornerstone of most immunosuppressive treatment protocols after solid organ transplantation in the United ...

Apical Resection Mouse Model to Study Early Mammalian Heart Regeneration

Cardiovascular disease plagues the whole world due to intensive lifestyle changes. Heart regeneration holds great promise for repairing and restoring cardiomyocytes lost due to injury and disease. In contrast to the robust cardiac regeneration of certain lower vertebrates, adult mammalian hearts typically show minimal capacity for heart regeneration and repair. However, recent studies have sparked considerable scientific interest with the finding that, between postnatal day 1 to 7 (P1 to P7), the neonatal mouse heart retains significant regenerative potential after apical resection (i.e., surgical amputation and exposure of left ventricular apex). One major controversy over this finding might be due to the diverse surgery-related procedures used in efforts to replicate or expand upon this important finding. These instructions dynamically present the materials and methodology for apical resection in a mouse model. The salient steps of this rodent survival surgery involve hypothermia anesthesia, thoracotomy, surgical amputation of heart ventricular apex, and suture and recovery of mice. The approach described could expand the application of the apical resection mouse model for cardiovascular research.

A step-by-step video protocol of apical resection is demonstrated in this study. Apical resection is a recently highlighted surgical approach in mammalian heart regeneration research. This study may promote the application of apical resection as a standard methodology in research into the mechanism underlying heart regeneration.

Cardiovascular disease plagues the whole world due to intensive lifestyle changes. Heart regeneration holds great promise for repairing and restoring ...

Novel Approach for Simultaneous Recording of Renal Sympathetic Nerve Activity and Blood Pressure with Intravenous Infusion in Conscious, Unrestrained Mice.

Renal sympathetic nerves contribute significantly to both physiological and pathophysiological phenomena. Evaluating renal sympathetic nerve activity (RSNA) is of great interest in many areas of research such as chronic kidney disease, hypertension, heart failure, diabetes and obesity. Unequivocal assessment of the role of the sympathetic nervous system is thus imperative for proper interpretation of experimental results and understanding of disease processes. RSNA has been traditionally measured in anesthetized rodents, including mice. However, mice usually exhibit very low systemic blood pressure and hemodynamic instability for several hours during anesthesia and surgery. Meaningful interpretation of RSNA is confounded by this non-physiological state, given the intimate relationship between sympathetic nervous tone and cardiovascular status. To address this limitation of traditional approaches, we developed a new method for measuring RSNA in conscious, freely-moving mice. Mice were chronically instrumented with radio-telemeters for continuous monitoring of blood pressure as well as a jugular venous infusion catheter and custom-designed bipolar electrode for direct recording of RSNA. Following a 48-72 hour recovery period, survival rate was 100% and all mice behaved normally. At this time-point, RSNA was successfully recorded in 80% of mice, with viable signals acquired up to 4 and 5 days post-surgery in 70% and 50% of mice, respectively. Physiological blood pressures were recorded in all mice (116&plusmn;2 mmHg; n=10). Recorded RSNA increased with eating and grooming, as well-established in the literature. Furthermore, RSNA was validated by ganglionic blockade and modulation of blood pressure with pharmacological agents. Herein, an effective and manageable method for clear recording of RSNA in conscious, freely-moving mice is described.

Anesthetized mice exhibit non-physiological systemic blood pressure, which precludes meaningful assessment of autonomic tone given the intimate relationship between blood pressure and the autonomic nervous system. Thus, a novel method to simultaneously record renal sympathetic nerve activity and blood pressure with intravenous infusion in conscious mice is outlined.

Renal sympathetic nerves contribute significantly to both physiological and pathophysiological phenomena. Evaluating renal sympathetic nerve activity ...

Shunt Surgery, Right Heart Catheterization, and Vascular Morphometry in a Rat Model for Flow-induced Pulmonary Arterial Hypertension

In this protocol, PAH is induced by combining a 60 mg/kg monocrotalin (MCT) injection with increased pulmonary blood flow through an aorto-caval shunt (MCT+Flow). The shunt is created by inserting an 18-G needle from the abdominal aorta into the adjacent caval vein. Increased pulmonary flow has been demonstrated as an essential trigger for a severe form of PAH with distinct phases of disease progression, characterized by early medial hypertrophy followed by neointimal lesions and the progressive occlusion of the small pulmonary vessels. To measure the right heart and pulmonary hemodynamics in this model, right heart catheterization is performed by inserting a rigid cannula containing a flexible ball-tip catheter via the right jugular vein into the right ventricle. The catheter is then advanced into the main and the more distal pulmonary arteries. The histopathology of the pulmonary vasculature is assessed qualitatively, by scoring the pre- and intra-acinar vessels on the degree of muscularization and the presence of a neointima, and quantitatively, by measuring the wall thickness, the wall-lumen ratios, and the occlusion score.

This protocol describes a surgical procedure to create a model for flow-induced pulmonary arterial hypertension (PAH) in rats and the procedures to analyze the principle hemodynamic and histological end-points in this model.

In this protocol, PAH is induced by combining a 60 mg/kg monocrotalin (MCT) injection with increased pulmonary blood flow through an aorto-caval shunt ...

A Chronic Cardiac Ischemia Model in Swine Using an Ameroid Constrictor

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but regardless of the approach, an in vivo model is needed to test each treatment. The pig is one of the most used large animal models for cardiovascular disease. Its heart is very similar in anatomy and function to that of a human. The ameroid placement technique creates an ischemic area of the heart, which has many useful applications in studying myocardial infarction. This model has been used for surgical research, pharmaceutical studies, imaging techniques, and cell therapies.
There are several ways of inducing an ischemic area in the heart. Each has its advantages and disadvantages, but the placement of an ameroid constrictor remains the most widely used technique. The main advantages to using the ameroid are its prevalence in existing research, its availability in various sizes to accommodate the anatomy and size of the vessel to be constricted, the surgery is a relatively simple procedure, and the post-operative monitoring is minimal, since there are no external devices to maintain. This paper provides a detailed overview of the proper technique for the placement of the ameroid constrictor.

The purpose of this protocol is to demonstrate the placement of a delayed constricting device (an ameroid constrictor) around a coronary artery in a swine model. This device creates an ischemic area of the heart that is useful for studying new diagnostic imaging techniques and new methods of treatment.

Cardiovascular disease remains the number one cause of mortality in the United States. There are numerous approaches to treating these diseases, but ...

Routine Screening Method for Microparticles in Platelet Transfusions

Platelet inventory management based on screening microparticle content in platelet concentrates is a new quality improvement initiative for hospital blood banks. Cells fragment off microparticles (MP) when they are stressed. Blood and blood components may contain cellular fragments from a variety of cells, most notably from activated platelets. When performing their roles as innate immune cells and major players in coagulation and hemostasis, platelets change shape and generate microparticles. With dynamic light scattering (DLS)-based microparticle detection, it is possible to differentiate activated (high microparticle) from non-activated (low microparticle) platelets in transfusions, and optimize the use of this scarce blood product. Previous research suggests that providing non-activated platelets for prophylactic use in hematology-oncology patients could reduce their risk of becoming refractory and improve patient care. The goal of this screening method is to routinely differentiate activated from non-activated platelets. The method described here outlines the steps to be performed for routine platelet inventory management in a hospital blood bank: obtaining a sample from a platelet transfusion, loading the sample into the capillary for DLS measurement, performing the DLS test to identify microparticles, and using the reported microparticle content to identify activated platelets.

Platelet inventory management based on screening microparticle content in platelet concentrates is a new quality improvement initiative in hospital blood banks. The goal is to differentiate activated from non-activated platelets to optimize the use of platelets. Providing non-activated platelets to hematology-oncology patients might reduce their high risk to become refractory.

Platelet inventory management based on screening microparticle content in platelet concentrates is a new quality improvement initiative for hospital blood ...

Simultaneous Electrocardiography Recording and Invasive Blood Pressure Measurement in Rats

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. Research focusing on cardiovascular disease models, potential cardiovascular therapeutic targets or pharmaceutical agents requires assessment of systemic arterial pressure and heart rhythm changes. In situations where radio telemetry systems are not available or affordable, the technique of femoral artery cannulation is an alternative way to obtain intra-arterial pressure waveform recordings and systemic BP measurements. This technique is economical and can be performed with standard equipment in animal facilities. However, invasive arterial pressure recording requires cannulation of small arteries, which can be a challenging surgical skill. Here, we present step-by-step protocols for femoral artery cannulation procedures. Key procedures include the calibration of the data acquisition system, tissue dissection and femoral artery cannulation, and setup of the arterial cannulation system for pressure recording. Surface electrocardiography recording procedures are also included. We also present examples of BP recordings from normotensive and hypertensive rats. This protocol allows reliable direct recordings of systemic BP with simultaneous electrocardiography.

Here, we describe a setup for simultaneous recording of electrocardiography and intra-arterial blood pressure (BP) in experimental rats, which can be done with standard equipment in animal facilities and can be applied to physiological or pharmacological studies to investigate pathogenic or therapeutic mechanisms in cardiovascular medicine.

For studies related to cardiovascular physiology or pathophysiology, blood pressure (BP) and electrocardiography are basic observational parameters. ...

A Rabbit Venous Interposition Model Mimicking Revascularization Surgery using Vein Grafts to Assess Intimal Hyperplasia under Arterial Blood Pressure

Although vein grafts have been commonly used as autologous grafts in revascularization surgeries for ischemic diseases, the long-term patency remains poor because of the acceleration of intimal hyperplasia due to the exposure to arterial blood pressure. The present protocol is designed for the establishment of experimental venous intimal hyperplasia by interposing rabbit jugular veins to the ipsilateral carotid arteries. The protocol does not require surgical procedures deep in the body trunk and the extent of the incision is limited, which is less invasive for the animals, allowing long-term observation after implantation. This simple procedure enables researchers to investigate strategies to attenuate the progression of intimal hyperplasia of the implanted vein grafts. Using this protocol, we reported the effects transduction of microRNA-145 (miR-145), which is known to control the phenotype of vascular smooth muscle cells (VSMCs) from the proliferative to the contractile state, into harvested vein grafts. We confirmed the attenuation of intimal hyperplasia of vein grafts by transducing miR-145 before implantation surgery through the phenotype change of the VSMCs. Here we report a less invasive experimental platform to investigate the strategies that can be used to attenuate intimal hyperplasia of vein grafts in revascularization surgeries.

The present protocol aims to experimentally create venous intimal hyperplasia by subjecting veins to arterial blood pressure for developing strategies to attenuate venous intimal hyperplasia following revascularization surgery using vein grafts.

Although vein grafts have been commonly used as autologous grafts in revascularization surgeries for ischemic diseases, the long-term patency remains poor ...

Implantation of Combined Telemetric ECG and Blood Pressure Transmitters to Determine Spontaneous Baroreflex Sensitivity in Conscious Mice

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. The baroreflex is a key homeostatic mechanism to counteract acute, short-term changes in arterial BP and to maintain BP in a relatively narrow physiological range. BP is sensed by baroreceptors located in the aortic arch and carotid sinus. When BP changes, signals are transmitted to the central nervous system and are then communicated to the parasympathetic and sympathetic branches of the autonomic nervous system to adjust HR. A rise in BP causes a reflex decrease in HR, a drop in BP causes a reflex increase in HR.
Baroreflex sensitivity (BRS) is the quantitative relationship between changes in arterial BP and corresponding changes in HR. Cardiovascular diseases are often associated with impaired baroreflex function. In various studies reduced BRS has been reported in e.g., heart failure, myocardial infarction, or coronary artery disease. 
Determination of BRS requires information from both BP and HR, which can be recorded simultaneously using telemetric devices. The surgical procedure is described beginning with the insertion of the pressure sensor into the left carotid artery and positioning of its tip in the aortic arch to monitor arterial pressure followed by the subcutaneous placement of the transmitter and ECG electrodes. We also describe postoperative intensive care and analgesic management. After a two-week period of post-surgery recovery long-term ECG and BP recordings are performed in conscious and unrestrained mice. Finally, we include examples of high-quality recordings and the analysis of spontaneous baroreceptor sensitivity using the sequence method.

The baroreflex is a heart-rate regulation mechanism by the autonomic nervous system in response to blood-pressure changes. We describe a surgical technique to implant telemetry transmitters for continuous and simultaneous measurement of electrocardiogram and blood pressure in mice. This can determine spontaneous baroreflex sensitivity, an important prognostic marker for cardiovascular disease.

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. ...