Equipment Needs & Protocol Considerations

www.transonic.com PV Equipment

Table of Contents

Transonic Scisense Pressure Systems & Catheters...... 3 Transonic Scisense PV Systems & Catheters...... 4 Research Equipment Sources...... 5 PV Catheter & ADV500 System Preparation...... 6 Balancing Pressure Sensors Before Use...... 7 Proper PV Catheter Placement in the Left Ventricle...... 8 Cleaning Guidelines for Catheters...... 12 Optimizing Catheter Life Span...... 14 Rodent Anesthesia Guidelines...... 17 Aseptic Surgical Guidelines...... 24 Surgical Recovery...... 25

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Transonic Scisense Pressure Systems & Catheters

Scisense Pressure Catheter compared to a reference trace and a fluid-filled pressure catheter at 300 BPM (5 Hz). The fluid-filled SP200 Pressure Measurement System catheter shows large distortion and wave artifact compared to the Scisense trace which reflects the true pressure waveform of the • Measure two channels of pressure simultaneously reference sensor. • Permits calculation of pressure gradients and pulse- wave velocity • Quick electronic two-point calibration • Compatible with all data acquisition systems (± 5V range required) 1.2F Mouse Pressure Catheter Solid-State Pressure Catheters • Single and Dual Pressure Sensor Catheters are available in all sizes ranging from 1.2F to 7.0F. • High frequency response ensures accurate detection of pressure waveforms and resulting calculations (dP/dt, Peak Pressure, MAP, etc.). • Variable placement of second pressure sensor on dual catheters is available to meet protocol needs. • Catheter tips can be customized to enable easy insertion. 5F Large Animal Pressure Catheter with pigtail tip • Smooth, flexible tubing allows easy insertion and navigation. *Note: Catheters not to scale

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Transonic Scisense PV Systems & Catheters

ADV500 Pressure-Volume Measurement System • Measure volume using either Conductance or Admittance modes • True volume in real-time using Admittance mode • Generate full hemodynamic reports and calculate measurements of contractility and stiffness • Two pressure inputs for pressure gradient or pulse- wave velocity Outer excitation electrodes create the electrical field which is detected by the inner sensing electrodes. • Compatible with all data acquisition systems (± 5V range required)

Pressure-Volume Catheters Pressure-Volume Catheters are available for all animal models. 1.2F and 1.9F Catheters are ideal for mouse and rat studies, 3.5F for rabbits, and 5.0F or 7.0F for all larger animals (canine, swine, bovine, etc.) • Excellent response at high frequencies such as rodent 1.9F Rat Pressure Volume Catheter & heart rates. 1.2F Mouse Pressure Volume Catheter • Smooth, flexible tubing allows easy insertion and navigation. • Catheter tips can be customized to enable easy insertion. 5F VSL Large Animal Pressure Catheter • Second pressure sensor available for 5.0F & 7.0F Catheters.

Variable Segment Length (VSL) Catheters VSL Catheters have 4 volume electrode options designed 7F VSL Large Animal Pressure Catheter to offer flexibility and ensure proper fit. VSL Catheters with pigtail tip are standard for all larger animal Catheters ranging from *Note: Catheters not to scale 3.5F - 7.0F, and optional for 1.2F and 1.9F sizes. Standard segment spacings are available for all Catheter sizes while custom designs can be made to order. Active segment can be changed at anytime during data collection.

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Research Equipment Sources

Additional equipment for surgical procedures may be acquired from the vendors listed below or from your vendor of choice.

Respirators Infusion Pumps Surgical Supplies CWE, Inc. Razel Scientific Instruments, Covidien (Kendall) www.cwe-inc.com Inc. Mansfield, MA Harvard Apparatus, Inc. Stamford, CT www.covidien.com Holliston, MA www.razelscientific.com/ www.kendallhq.com ww.harvardapparatus.com Rattus (Kent Scientific) BD Kent Scientific Corp. (Rattus) Torrington, CT Franklin Lakes, NJ Torrington, CT www.kentscientific.com or www.bd.com www.kentscientific.com or www.rattus.com Harvard Apparatus www.rattus.com Holliston, MA Sonomicrometers ww.harvardapparatus.com Micromanipulators / Stands Sonometrics Corporation Fine Science Tools, Inc. London, Ontario, Canada Cleaning / Disinfecting Agents Foster City, CA www.sonometrics.com Alconox Inc. www.finescience.com White Plains, NY Techni-Tool Inc. Fluid Infusion www.alconox.com Worcester, PA Instech Laboratories, Inc. Ruhof www.techni-tool.com Plymouth Meeting, PA Mineola, NY Stoelting Co. www.instechlabs.com www.ruhof.com Wood Dale, IL Advanced Sterilization www.stoeltingco.com Surgical Monitoring and Products (J & J) Heating Irvine, CA www.aspjj.com Langendorff Apparatus Indus Instruments Hugo Sachs Electronik: Webster, TX Harvard Apparatus www.indusinstruments.com Holliston, MA ww.harvardapparatus.com Rattus (Kent Scientific) Torrington, CT www.kentscientific.com or www.rattus.com Radnoti Glass Technology, Inc. Monrovia, CA www.radnoti.com

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PV Catheter & ADV500 System Preparation

Follow all directions supplied with the Pressure-Volume Catheters and ADV500 PV System including manuals and quick reference guides.

Catheter Preparation Pre-soak the tip of the PV Catheter in 0.9% saline for ~20 min before its insertion (Fig. 1). After soaking, the pressure senor(s) can be balanced using the [Course] and [Fine] adjustments on the ADV500. See “Balancing Pressure Sensors Before Use” on page 7 for recommended procedure.

ADV500 / ADVantage System Preparation Connect the ADV500/ ADVantage system to the data acquisition software, ensuring all channels are calibrated. For the Admittance method, constants for “Heart Type” (sigma-epsilon ratio), blood resistivity and stroke volume reference must be input prior to data collection. For Conductance mode, all volume calibration will be performed post-acquisition. • For “Heart Type,” choose from “Normal” or “Infarct,” or use a custom value

• For blood resistivity, use a known value or get a measurement using the Fig. 1: Soak PV Catheters in calibration probe. saline prior to use • Determine stroke volume (SV) by using cardiac MRI, CT scan, high resolution echocardiography, Swan-Ganz thermo-dilution catheter or aortic Flowprobe. It is recommended to determine SV independently for a small sample of the species to be studied and use the average value for subsequent experiments.

Calibration Probe: Determine Blood Resistivity If possible, before the procedure, collect ~2 ml of non-heparinized arterial blood to measure blood properties. Place the tip of Calibration Probe in the sample of freshly drawn blood (35-37°C) and touch the blood meniscus (Fig. 2). Press [OK] on ADV500 box to sample blood resistivity (ρ). Please do not submerge the Probe into the blood. The procedure has to be finished quickly as blood will change properties as it clots. If a resistivity measurement cannot be taken, a known value must be used instead. Fig. 2: Calibration Probe is used to determine blood resistivity.

When using electrocautery or defibrillator devices with Pressure or PV Catheters, ensure that the animal has been grounded and the Catheter unplugged from the control unit (can be left in place). Failure to take appropriate precautions to avoid transferring of electrical currents can cause damage to the Transonic Scisense Equipment or personal harm.

Fig. 3: ADV500 Pressure-Volume Measurement System

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Balancing Pressure Sensors Before Use

Scisense Pressure Catheters are built with piezo resistive strain gauges that detect pressure through a flexible rubber membrane. Due to the mechanical properties of the rubber and the nature of gauge pressure sensors, proper use requires an understanding of how to properly balance the sensor.

D

Atmospheric BC Pressure A Water Column Atmospheric Atmospheric 0.75 mmHg per cm Atmospheric Pressure Pressure Pressure Offset Artifact Zeroed Offset Artifact Zeroed Offset Artifact Sensor Sensor Sensor Sensor

Atmospheric Atmospheric Atmospheric Atmospheric Pressure Pressure Pressure Pressure

(A) Ideal Pressure Sensor referenced to atmospheric pressure. The forces on both sides of the sensing membrane balance so the output is zero. (B) In the real world, there is always a mechanical or electrical factor that is going to cause an imbalance across the pressure sensing membrane. The artifact will vary between catheters and associated amplifiers. (C) For this reason, each control box comes with an offset correction control which can be used to counter balance the offset artifact. This electronically zeros the output. (D) The Sensor can then be submerged in a beaker of water to a given depth. Since the artifact has been cancelled out and the atmospheric pressure is equal on both sides, the Sensor will output 0.75 mmHg for each centimeter of water depth it is submerged. Balancing your catheter before use: 1. Air Calibration: Calibrating the Sensor in air should provide the best result. There are, however, two considerations: a. Since the Catheter should have been soaked in fluid for some time, it will be wet when exposed to air. The exothermic effects of evaporation could exceed the temperature compensation features that are built into the Catheter. b. Since the Catheters are very sensitive, any motion might be detected when holding the Catheter. Combined with exothermic events, this can result in a wandering signal. If the software used to analyze the signal is in an auto-gain mode, the effect is exaggerated even further. 2. Calibrating the Sensor in saline: Body temperature saline used to soak the Catheter is the best environment for Catheter calibration. However, the user needs to be aware of the offset value that the saline will create. If the Catheter is zeroed under a 5 cm column of saline when it is removed from the water, the reading will be -3.5 mmHg. Inserting the Catheter into a ventricle with this offset would result in a negative EDV value. The best way to calibrate the Catheter is to hold it just under the surface (meniscus) of body temperature saline as it is being balanced. The offset should be minimal for a Catheter under a few mm of water. Any minor signal wandering as the Catheter is transferred to the blood vessel can be ignored as either motion or temperature artifact.

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Proper PV Catheter Placement in the Left Ventricle

Viewing the pressure vs. magnitude loops and phase signal during catheter positioning in the ventricle can assist in proper placement. Correct placement of the Catheter in the ventricle is important for accurate data collection. Always use care when inserting the Catheter past the aortic valve as excessive force can cause Catheter damage, especially in small, rodent Catheters. Variable Segment Length (VSL) Catheters have four segment length settings. Select the longest segment length that gives physiological shaped pressure vs magnitude loops. The typical range of magnitude values for healthy animals with the specified body weights and stroke volumes are as shown:

Stroke Magnitude │γ│ Phase Species Body Weight The phase signal allows you to volume (SV) Variation (degree) visualize the proximity of the Catheter Mouse 20 - 25 g 18 - 23 μL ≥ 200 μS 4 - 8 to the heart wall and can help in Rat 300 - 400 g 270 - 360 μL ≥ 500 μS 3 - 7 Catheter placement. The signal should Pig 35 - 45 kg 30 - 40 ml ≥ 2.5 mS 1 - 5 be periodic in shape with a relatively low mean value (< 10°). Position If using a VSL Catheter, start with the shortest segment length the Catheter for the lowest mean (segment 1) for initial insertion. phase signal (center of the LV). If the phase signal is excessively noisy, try repositioning the Catheter slightly or remove any sources of interference that may be attached to the animal.

Aortic valve

Proximal electrodes

Pressure sensor

Distal electrodes Four selectable segment lengths (active rings in red). Segment 1 is the shortest (for smaller hearts) while segment 4 is the longest (for larger hearts). The distal electrode pair (1 & 2) remain the same for all segments.

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Proper PV Catheter Placement in the LV Cont.

The evolution of pressure vs magnitude loops as the Catheter is inserted into the left ventricle from the aorta follows: 1. Pressure sensor is in the LV, just past the aortic valve. The two distal volume electrodes are in the LV. The two proximal electrodes are in the aorta.

2. Pressure sensor is in the LV, just past the aortic valve. The two distal volume electrodes are in the LV. The two proximal electrodes are in the aorta, very close to the aortic valve.

3. Pressure sensor is in the center of the LV. The two distal volume electrodes are in the LV. The two proximal electrodes are on either side of the aortic valve. • If the Catheter is too long for the ventricle size being studied, this may be the best position possible. If available, a smaller Catheter should be used instead.

4. Pressure sensor and volume rings are completely inside the LV. The most proximal electrode is situated very close to the aortic valve in the LV. • This is the ideal Catheter position. • Once the ideal depth has been determined, check the phase signal to ensure the Catheter is appropriately centered in the ventricle.

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Proper PV Catheter Placement in the LV Cont.

1. Catheter tip is very close to the apex. The distal electrodes almost touch the apex. • This Catheter position is acceptable. During IVC occlusions, the proximal electrodes could be pushed out of the LV into the aorta.

2. Catheter is jammed into the apex. The |Y| variation is low (≈ 100 μS) • Non-ideal position and must be avoided, if possible. This position causes premature ventricular contractions (PVC) in rats and large animals.

Once the Catheter is fully inserted, segment size can be adjusted on VSL Catheters. Increase the segment length one size at a time and observe the shape of the loops. Continue increasing the spacing until the shape of the loops no longer appears physiological. Then return to the previous segment. The following is an illustrative example of segment selection:

Segment 1 Active electrodes: 1, 2, 3, 4 • Selected segment is too short for the LV • The shape of the Pressure vs. Magnitude loops look physiological • Calculated volumes will be low due to missing volume at the base of the heart (near the valve) • Select segment 2

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Proper PV Catheter Placement in the LV Cont.

Segment 2 Active electrodes: 1, 2, 4, 5 • Selected segment is too short for the LV • The shape of the Pressure vs. Magnitude loops look physiological • Calculated volumes will be low due to missing volume at the base of the heart (near the valve) • Select segment 3

Segment 3 Active electrodes: 1, 2, 5, 6 • Selected segment is the ideal length for the LV • The shape of the Pressure vs. Magnitude loops look physiological • Calculated volumes will be accurate • Select segment 4

Segment 4 Active electrodes: 1, 2, 6, 7 • Selected segment is too long for the LV • The shape of the Pressure vs. Magnitude loops appears distorted • Calculated volumes will be inaccurate • Select segment 3 since it is the appropriate segment length for the LV

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Cleaning Guidelines for Catheters

Always clean Catheters immediately after use. Failure to properly and Promptly clean Catheters may cause sensor failure. Do not use Ultrasonic Cleaners for the catheters.

Follow all directions for the cleaning and disinfecting agents. 1. Immediately after use, immerse the Catheter in distilled water or saline for approximately 5 minutes. 2. Soak the Catheter in an enzymatic cleaning solution to remove all traces of biological material (i.e. Tergazyme by Alconox Inc). This should take between 0.5 and 2 hours depending on cleaning agent and level of soil on the Catheter. 3. Optional: Immerse Catheter in disinfecting agent (i.e. Cidex® by Rinse Catheter in distilled water or saline (Steps 1 & 4) ASP) to remove all traces of viable microbes. Do not use glutaraldehyde solutions containing surfactants such as Cidex® 7 or Cidex® Plus, or solutions containing hydrogen peroxide like Sporox. Do not use Cidex® PA. 4. Rinse the Catheter by soaking in distilled water for 1 - 5 minutes to remove all traces of cleaning agents . Immerse Catheter in cleaning agents (Steps 2, 3 & 7) 5. Dry the Catheter by gently wiping or placing on a paper towel or gauze. Do not air dry or use alternative drying methods. Never apply direct pressure to the pressure sensor membrane. 6. Before returning the Pressure Catheter to its original packaging for storage, use magnifications to check for blood or tissue residue on the Catheter tip. If any is found, repeat the cleaning process. 7. Optional: Immerse Catheter in an acid-based cleaner to remove metal oxides from the ring electrodes on Pressure-Volume Catheter in storage foam (Step 8) Catheters for 1 -2 minutes (i.e. Citranox® by Alxonox Inc). 8. Store the dry Catheter in its original packaging. Position the pressure sensor within the foam cutout to prevent damage. Single-use Catheters should be disposed of after use, according to local waste disposal regulations.

Effect of catheter cleaning on performance Proper Catheter cleaning ensures proper performance of pressure and pressure-volume systems and the ability to confidently present good data. Catheter cleaning is an important task and requires specific attention after each procedure. Soiled Catheters are not able to perform at their 1.2F Rodent Pressure Volume Catheter before (top) and after best, even if the best position is found in the heart cavity. It (bottom) proper cleaning. A 30 min wash in 1% Tergezyme is important to visually inspect Catheters under a microscope (Alconox Inc.) removes the dried liquid and blood. after cleaning to ensure that all contaminates have been removed. Particularly stubborn soil may require additional cleaning cycles to fully remove.

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Cleaning Guidelines for Catheters Cont.

Effect of catheter cleaning on performance Cont. It is important to note that there are many other factors, besides Catheter cleanliness, which impact performance. See “Optimizing Catheter Life Span” on page 14 for more information on Catheter use best practices. Additionally, improper PV Catheter positioning within the ventricle can produce anomalous data. Follow the instructions in “Proper PV Catheter Placement in the Left Ventricle” on page 8 for ideal Catheter placement.

Data from a Catheter before proper cleaning (left) and after proper cleaning (right) in the LV of a mouse. Note the degraded quality of the phase signal with the dirty Catheter which shows increased variability and lacks the ideal sinusoidal shape seen with the clean Catheter. The magnitude and dP/dt graphs both shows damping from the dirty Catheter, see table below for values. Additionally, left ventricle pressure signal can be dampened by tissue or dried liquid present on the Catheter, constricting the sensor to the point where it damages the pressure sensing surface inside.

PV Parameters Example Dirty Catheter Values Clean (Expected) Catheter Values Pressure (mmHg) systolic: < 90 diastolic: < 6 systolic: 90-120 diastolic: 1-6 Phase (degrees) 0 - 10 and above 4 - 8 Magnitude (uS) high to low variation < 200 high to low variation > 200 dP/dt max (mmHg/s) < 4000 6000 - 10000, up to 17000 Mouse under 1.5% isoflurane anesthesia with heart rate between 500-600 bpm

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Optimizing Catheter Life Span

Catheters are delicate measurement devices: always handle with care. Failure to properly handle catheters may result in voiding the warranty. Follow all instructions in the Quick Start Guides, Inserts and Operator’s Manuals.

In order to get the longest life possible from a Scisense Pressure or Pressure-Volume Catheter, it is important that the user have some appreciation of the delicate nature of Catheters. The sensing surfaces, tube wall and conducting wires are measured on the order of microns. However, when appropriate precautionary measures are taken, Catheters can be successfully reused for many experimental protocols.

• Damage to the Catheter shaft from excessive force is the most common cause of Catheter failure. While the shaft material is very strong, it has a yield point, and will break if it is deformed past this point. The catheter shaft will also be weakened by any micro cuts or abrasion that may be inflicted during the course of its use. Correct Catheter handling with fingers. Grip is along • Before handling a functional Catheter, practice catheter the shaft, well behind the sensitive Catheter tip. handling and insertion with a dummy Catheter. • If you chose to handle the Catheter with forceps, place small pieces of polyethylene (PE) tubing over the forceps’ tips. This will protect the Catheter body against kinks or abrasions from the sharp forceps’ edges. When using forceps to handle the Catheter, please be aware that the forceps or grasping ends are not designed to manipulate such a delicate shaft. • When starting to work with Catheters, please use a surgical microscope to estimate and coordinate the actual hand grip and applied strength under a variety of magnifications. This will help you with Catheter/hand coordination and help determine the amount of force required to hold the Catheter. • Ensure that you are not grasping the Catheter with either fingers or forceps close to the area where metal rings or Place polyethylene tubing over the forceps’ tips to pressure sensor(s) are located. Applying pressure directly to the protect the Catheter body from damage. pressure sensor or metal rings can cause significant damage. • Even when using protected forceps, the user must be aware of the force generated on the Catheter shaft. It should not be necessary to exert force to any extent that noticeably deforms the diameter of the Catheter. Crushing the tube flat will destroy the strength and possible damage the wires inside. Note: This type of damage is easy to identify and will not be covered by the warranty.

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Optimizing Catheter Life Span Cont.

• Be gentle when inserting and withdrawing Catheters, especially when navigating past tie off sutures and heart valves, as this is the second most common way for Catheters to sustain damage. • Consider using inhalation anesthesia to anesthetize the animal to ensure complete control over body position for the entire duration of the catheterization (from insertion to withdrawal). Any sudden change of position from an uncontrolled animal or anesthesia replenishment might damage (kink) the Catheter. • When catheterizing a vessel, ensure the area where the Catheter is inserted is well surgically prepared and maintained without an excessive amount of vascular sheets (adventitia). When placing a Catheter in the ventricle or atria ensure that the pericardium is removed close to insertion site and that the appropriate sized needle is used to puncture the tissue. Never try to force the Catheter through a thick layer of vascular adventitia or cardiac tissue.

• If using a carotid artery access, the animal is usually in Correct Catheter handling with forceps. PE tubing covers the supine position, ventilated with 1-2% of Isoflurane, and all forceps’ tips and the grip is along the shaft, well behind the nociceptive withdrawal-reflexes are deficient. Insertion is sensitive Catheter tip. through a well-cleaned segment of common carotid artery. Complete preparation and surgery can be seen on our website www.transonic.com. Access through the right carotid artery works better than the left carotid artery in LV catheterization (1). • The suture placed around the common carotid area and tied over the segment during surgery has to be positioned such that the sensing surface of the Catheter does not bear direct contact with the suture. Moreover, it is important that holding and supporting sutures are not tied to the point where the Catheter requires a strong force to pass through (during either insertion and withdrawal). Be careful, when tying off sutures, to stabilize the Catheter during measurements. If the sutures are tied too tight, they can damage the Catheter shaft. Incorrect Catheter handling with forceps. • When passing the Catheter into the ascending aorta and through the aortic Note how the grip is on the sensitive Catheter tip. valve to enter the left ventricle (LV), the sensor tip of the Catheter often encounters resistance at the valve entrance. If you are using a ventilation set-up, long supine-axis position can be adjusted to accommodate the Catheter in such way that the Catheter passes more in line with the supine- axis of the animal. This manoeuvre can be achieved by pulling on the front paws to reposition the animal, while slowly withdrawing and inserting the Catheter without an excessive force. Trying to force the Catheter past this point might result in the Catheter bending too much and inducing a permanent crimp in the Catheter shaft.

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Optimizing Catheter Life Span Cont.

• Failure to properly clean the Catheter post- intervention is detrimental to its performance and lifespan. See “Cleaning Guidelines for Catheters” for detailed instructions. • Tissue left on the Catheter can actually constrict the sensor to the point where it damages the pressure sensing surface inside. • Tissue left on the Catheter will eventually alter the properties of the sensor interface. While it will not stop the sensor from working, it will change the baseline offset and potentially alter sensitivity. • The bulk of cleaning should be done with an enzymatic cleaner that will dissolve any tissue. If tissue is observed to be stuck on the sensing surface of the Catheter, the tissue should be 1.2F Rodent Pressure Volume Catheter before (top) and after (bottom) removed with extreme care under a microscope. proper cleaning. A 30 min wash in 1% Tergezyme (Alconox Inc.) removes • Catheters should never be pulled through a the dried liquid and blood. cleaning cloth or “wiped” in such a manner where the cloth puts force on the sensor surface. • Cleaning your Catheter and looking for any residual material under a microscope is a good opportunity to look for damage from use. If shaft roughness or marks from forceps/tweezers are observed, it is a good indication that the surgical procedure needs to be adjusted. There should be no visible wear or tear on a properly handled Catheter shaft.

Reference (1) Migneco F, et. al. “New and simplified method for multiple left ventricle catheterizations in small animals.” Interact CardioVasc Thorac Surg 2008; 7: 925-927.

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Rodent Anesthesia Guidelines

There is no single best choice for anesthetic agents as procedure, parameters of interest, and animal type all impact anesthesia choice. Always check what is currently available and allowed with your Institutional Animal Care & Animal Use Committee and make sure that the anesthetic agent is balanced with proper analgesics. It is important to note that the availability of anesthetic agents changes and is dependant on your institution and country.

Considerations Related to the Procedure Considerations Related to the Drug(s) Used • Type of procedure • Drug safety and ease of use • Projected length of the procedure • Appropriateness for the procedure including • Amount and type of pain/distress anticipated administration method • Study goals (are important parameters influenced by • Appropriateness for the animal certain drugs?) • Side effects • Survival or terminal study (agents associated • Equipment and training required for safe use with prolonged recovery or delayed effects may • Previous experience using the agent(s) be approved for terminal studies while deemed inappropriate for survival procedures) • Cost and status as controlled or uncontrolled drug • Acute or chronic study Summary: Anesthetic Agents Should Considerations Related to the Animal • Provide an appropriate depth and length of anesthesia and analgesia without affecting • Species and strain important study parameters • General condition and underlying health problems • Be appropriate for the animal given its species, • Age medical history and physical condition • Sex • Have minimal side effects • Weight • Be safe for both the animal and the personnel • Previous Drug Exposure administering anesthesia • Nutritional Status • Time of day as related to circadian rhythm • Ability to maintain body temperature (preventing hypothermia due to heat loss) • Numbers of animals to be anesthetized simultaneously Note: There can be remarkable variation in response to anesthesia. Investigators should monitor anesthesia closely in each animal and make appropriate modifications in the anesthetic regimen when necessary

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Rodent Anesthesia Guidelines Cont.

Cardiovascular Effects of Anesthetics Many common anesthetics have a significant effect on cardiovascular measurements and can obscure or confound study results; sometimes over a longer period of time than anticipated. It is therefore necessary to choose an anesthesia protocol with care. For the purpose of cardiac experimental procedures general anesthesia is recommended, however dissociative anesthetics in combination with a sedative agent may be used as well. During the experimental procedure, management of anesthesia has to be catered to any underlying or experimentally caused cardiovascular disorder. For example, experimentally induced aortic stenosis (trans- aortic banding or constriction) requires anesthesia which avoids systemic vasodilation and tachycardia while preserving sinus rhythm such as a synthetic narcotic based anesthesia.

Inhaled (Halogenated Ether) Anesthetics It is known that inhaled anesthetics may cause circulatory depression at concentrations required to produce general anesthesia. In addition, each individual inhalation anesthetic has selective dose-dependent effects on cardiovascular function (sympathetic reflexes, intravascular volume status, vascular smooth muscle tone, myocyte contraction and relaxation, acid-base status etc.). For this reason, circulatory interactions of inhaled anesthetics might limit the anesthetic dose. Consequently, some laboratories combine inhaled anesthetics with sedatives or hypnotics to produce the necessary general anesthesia. Others empirically developed state of the art mono-anesthetic protocols using minimum amount of inhalation anesthetics to mimic close to fully- conscious state while collecting data. Drop of blood pressure (BP) caused by inhalation anesthetics is a direct result of dose-dependent vasodilation accompanied by an afterload reduction and depression of myocardial contractility and an indirect result of attenuation of sympathetic nervous system. Decrease in BP during Isoflurane induced general anesthesia is so predictable that some laboratories often use this as a sign for assessing the depth of anesthesia. Halogenated anesthetics decrease global LV systolic function at any given LV loading condition or at any given degree of underlying sympathetic tone. Experimental studies suggest that these agents cause minimal changes in LV diastolic compliance but impair LV diastolic relaxation in a dose-dependent manner. These agents have minimal direct effects on LV preload, but rather EDP may increase during anesthesia because of impaired diastolic filling and decreased cardiac output (CO). The administration of inhaled anesthetics to experimental animals with cardiovascular diseases has some advantages. Most inhaled anesthetics are myocardial depressants with negative inotropic properties which decrease contractility and thus decrease myocardial oxygen demand. Arterial vasodilation combined with preserved coronary perfusion maintains oxygen delivery to the heart. Adequate oxygen delivery combined with a decreased demand for oxygen creates a more favorable myocardial oxygen balance in hearts with coronary insufficiency. Additionally, the vasodilating and antihypertensive actions of inhaled anesthetics effectively control an increase in BP in response to surgical pain. Inhalation anesthetics have a proportionally greater negative inotropic effect on diseased myocardium compared with normal myocardium. In the case of an experimentally induced septic shock by injection of LPS or cecal puncture, profound ventricular dysfunction may not tolerate the cardiovascular depressant effects of inhaled anesthetics given in concentrations that are needed to produce the anesthesia. The pro-thrombotic side effect of sepsis causes decreased coronary perfusion pressure which prevents adequate oxygen extraction via Fick’s principle. In this case cardiac oxygen demand excesses the rate of consumptions (MVO2) causing a negative oxygen balance which further depresses cardiac function.

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Rodent Anesthesia Guidelines Cont.

Rodent Anesthesia Breathing Circuitry : Open System is the traditional method of dipping ether or chloroform on gauze, later modernized by the Schimmelbusch mask and used until about 1950. Semi-open System is commonly used today and includes all the Mapleson systems. This is typically used for animal anesthesia induction, usually a single branched system that uses a valve to control the pressure of the gas, and allows for waste gas to leave the system. This system can be further characterized by high fresh gas inflow in order to stop re-breathing of expired CO2.

Semi-closed and Closed Systems use a CO2 absorbent and thus gases are re-circulated; the classification (semi- open vs closed) is defined by the amount of fresh gas flow. These systems are mainly used for maintenance of anesthesia following induction. Additionally, they can be used for anesthesia induction, but this is a slower process than using a semi-open system.

Expired gases from the animal pass through a container in the breathing system which contains a CO2 absorbent to remove CO2 from the expired gases. This method requires a high level of animal monitoring, especially levels of inspired and expired CO2 and the anesthetic agent. This absorbent, by an exothermic chemical reaction removes the CO2, thus allowing an animal’s expired gases to be re-breathed. Because of this exothermic chemical reaction, some warmth and humidity is added to the inspired gases. In this setting, the animal’s expired gases are recirculated, allowing for a reduced inflow rate of additional fresh gas. Breathing system components: 1. Fresh gas intake Fresh gas intake Fresh gas intake (O2, medicinal air etc.) 2. Adjustable pressure and/or volume Flow control Flow control limiting valve 3. Connection to animal Vaporiser Vaporiser (ventilator) 4. Waste gas connection tubing or Isoflurane 0-5% Isoflurane 0-5% Filter CO 2 canister or anesthesia gas absorber absorber active gas or gas evacuation evacuation system system Ventilator

Semi-open system Semi-closed system Schema of Isoflurane inhalation semi-open circuit Schema of Isoflurane inhalation semi-closed or closed (gases are not recirculated). Unidirectional valves system for rodent maintenance anesthesia. Unidirectional permit pressure driven flow through the vaporizer to valves permit pressure driven flow through the vaporizer the Anesthetizing Box; exhaled gases are routed into into the inspiratory limb of circle system. Exhaled gases Filter canister (removal of excess of Halogenated are routed into expiratory limb and recirculated through gases) or into active gas evacuation system. use of CO2 absorber. A bidirectional valve positioned in the expiratory limb permits gases to be evacuated if needed (e.g. high pressure develops).

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Rodent Anesthesia Guidelines Cont.

Sedatives-Hypnotics This group of anesthetics include barbiturates, benzodiazepines, etomidate, propofol and ketamine. They are used for pre-surgical sedation, producing immediate loss of consciousness, to supplement the actions of the inhaled anesthetics, and to provide sedation in the immediate postoperative period. The circulatory effects of individual agents are an important consideration for subjects with CV disease. The sedative-hypnotics have direct effects on cardiac contractility and vascular tone in addition to indirect effects on autonomic tone. Barbiturates (e.g. sodium pentobarbital, thiopental and methohexital) are anxiolytics, hypnotics, anticonvulsants and weak analgesics with negative inotropic effects. They produce dose-dependent decrease in dP/dt and the force-velocity relationship of ventricular muscle. Induction of general anesthesia with barbiturates is associated with a decrease in blood pressure (BP), heart rate (HR) and cardiac output (CO). In comparison with barbiturates, propofol appears to cause less myocardial depression. Mean arterial pressure (MAP) decrease after propofol is attributed primarily to both arterial and venous dilatation. Propofol is well suited for continuous i.v. infusion for sedation because it has a short duration of action and can be titrated to effect. Propofol is usually combined with opioids (Fentanyl, Sufentanyl etc.) for its lack of analgesia. Etomidate and ketamine are administrated for rapid induction of general anesthesia in experimental animals with pre-existing hemodynamic compromise because they generally cause little or no change in circulatory parameters. Etomidate has virtually no effect on myocardial contractility even in diseased ventricular muscle. For its endocrine and neuroendocrine non-anesthetic interferences it is limited to short-term use as an i.v. induction agent. Ketamine often increases HR and BP and causes bronchodilation because of its sympathomimetic properties. Ketamine has other beneficial effects including analgesia, anesthesia, and direct negative inotropic and vasodilatation effects

Narcotics (opioid) anesthetics Narcotic-based anesthetics offer the advantages of profound Advantages of inhalation anesthesia as analgesia, attenuation of sympathetically mediated cardiovascular reflexes in response to pain, and have compared with injectable anesthetics virtually no direct effects on myocardial contractility. Even • Easily controllable cardiovascular depression though narcotics have little direct action on the heart, they • Reduced impact on liver functions may cause profound hemodynamic changes indirectly by attenuating sympathetic nervous tone while decreasing • Reduced impact on kidney functions serum catecholamine levels, which may cause indirect cardiac • Encourages rapid recovery depression. • Allows superb control while on anesthesia In addition, other inconveniences encountered with narcotic- • Easy maintenance of surgical anesthetic based anesthetics include difficulty estimating required depth dose, predicting the duration of postoperative narcotic- induced respiratory depression, and ensuring hypnosis during • Dose and volume can be easily adjusted operation. Rapid administration of narcotics (Fentanyl) is also • Less stress on subject as compared to associated with muscle rigidity of the thoracic and abdominal injections musculature that may impede the ability to ventilate the • More predictable pharmacokinetics patient immediately after the induction of general anesthesia. Development of short-acting narcotic anesthetics may improve the ability to control anesthetic depth without prolonging recovery time. Ultra-short-acting narcotics (Remifentanyl) may have a unique niche in cardiac anesthesia because their effect is terminated immediately on stopping the drug infusion due to rapid in vivo ester hydrolysis.

PV Workbook RPV-3-wb Rev A 2014 Technical Note: RL-67-tn 20 PV Equipment

Rodent Anesthesia Guidelines Cont.

Category Agent Species & Dose (mg/kg) Route* Hemodynamic effects* PMID Citation Alphaxolone (Alfaxan) Mice: 15 IV increased HR, decreased MAP 17319964 Alphaxolone/Alphadolone 11098097, Rats: 18/6 IP vasodilation (Saffan) 11575348 Chloral hydrate Rats: 300 - 400 IP minimal cardiopulmonary depression 8355479 Alpha-chloralose1 Rats: 50-55 IP minimal cardiopulmonary depression 19003937 Fentanyl/Droperidol (Innovar-Vet) Mice: 0.078/3.9 IM vasodilation 15288130 decrease HR, SV & CO, Fentanyl/Medetomidine Rats: 0.3/0.3 IP 22561119 cardiorespiratory depression Mice: 75/0.2/1 Vasodilation, cardiorespiratory 19001064, Propofol/Fentanyl/Medetomidine IP Rats: 100/0.1/0.1 depression 20819392 Vasodilation, cardiorespiratory Propofol/Remifentanyl Mice: 50-200/0.2-1 IP 17640460 depression Ketamine 80- 200 IM good HR & BP 18172330 Ketamine/Diazepam (Valium) Mice: 100/5 Rats: 40/5 IP minor cardiorespiratory depression 7278122 Mice: 80-150/7.5-16 cardiorespiratory depression (MAP & 15155266, Ketamine/Xylazine (Rompun) IP, IM Rats: 40-80/5-10 CO), arrhythmia 7278122 Mice: 50-75/1-10 Anesthetics - Ketamine/Midazolam IP decreased MAP & CO 16174120 Injectable Rats: 60/0.4 Ketamine/Acepromazine Mice: 100/5 IP minor CV depression, hypotension 23382271 Mice: 100/2.5/2.5 Ketamine/Xylazine/Acepromazine IP, IM good MAP & HR 11924805 Rats: 40/8/4 decreased CO, MAP & HR; increased 15155266, Pentobarbital Na (Nembutal2) Mice: 30-90 Rats: 30-60 IV, IP ESV & EDV 15027618 good CI, minor cardiorespiratory Tiletamine/Zolazepam (Telazol) Rats: 20 - 40 IM, IP 17343357 depression cardiorespiratory depression, Thiamylal (SuritalR) Rate: 25 - 50 IV, IP 1637605 arrhythmia Cardiorespiratory depression, Thiopental Na (PentothalR) Mice: 30-40 IV, IP 18172330 decreased BP 7278119, Etomidate Mice: 22-25 IP decreased HR, good CO & MAP 12814659 Urethane1 Mice: 800 - 1300 IP good MAP, & CO 15155266 Urethane/Etomidate/Morphine1 Mice: 750/20-25/1-2 IP good MAP & CO 15604134 moderate cardiopulmonary Tribromoethanol1 (TBE or Avertin) Mice: 250 Rats: 150 IP 16884172 depression Mice: 0.1-1.5% 18550865, Isoflurane (Forane) Rats: 0.25-2.5% in pure Inhalation Vasodilation, decreased BP, good CO 12003817, O maintenance 22492676 Anesthetics - 2 Inhalant3 Desflurane To effect (4-6%) Inhalation Vasodilation, decreased BP 22929732 Rats: 3.5-4% in pure O 21778336, Sevoflurane 2 Inhalation Vasodilation, decreased BP maintenance 22167771 *SC=subcutaneous, IM= intramuscular, IP= intraperitoneal, PO=orally, IV=intravenous, HR = heart rate, SV = stroke volume, MAP= mean arterial pressure, CO= cardiac output, CI= cardiac index, ESV = end systolic volume, EDV = end diastolic volume, 1. Terminal Studies only. 2. Dilute stock solution to accurately dose animals 3. These agents should be used only in ways that prevent exposure to personnel. Induce anesthesia in a closed container and maintain with a nose cone in an appropriately ventilated hood.

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Rodent Anesthesia Guidelines Cont.

Anesthesia Tips & Considerations Category Agent Dose (mg/kg) Frequency Route Both: Both: once • When anesthetizing post-MI animals, Atropine SC 0.02-0.05 at induction maintenance of coronary artery pressure helps Anticholinergics Both: Both: once limit tachycardia. Glycopyrrolate SC 0.01-0.02 at induction • Induction of anesthesia can cause arrhythmias Rats:100-300 Rats: 4 hrs Acetaminophen PO (junctional rhythms). Treat by reducing the dose Mice: 300 Mice: daily of inhalation anesthetic or administering an Aspirin Both: 100 Both: 4 hrs PO anticholinergic. Analgesic (NSAID) Rats:12 hrs Carprofen Both: 5 SC • Halogenated volatile inhalation anesthetics Mice: daily (isoflurane) should be used in a vented hood to Flunixin Both: 1.1-2.5 Both: 12 hrs SC, IM reduce operator exposure during procedures. Ibuprofen Both: 7.5 Both: daily PO • It is advisable to monitor blood gases before, Rats: 0.05-2 Both: Butorphanol SC during and after anesthesia to ensure normal Mice: 0.05-5.4 2-4 hrs metabolism and prevent the development of Analgesic Meperidine Both: 10 - 20 Both: 2-3 hrs SC, IM (Opiate) alkalosis or acidosis. Morphine Both: 10 Both: 2-4 hrs SC, IM • Anesthetized animals do not completely close Pentazocine Both: 10 Both: 3-4 hrs SC their eyelids. Therefore, they are at risk of corneal desiccation and ulceration. It is advisable to Body Temperature protect their eyes with sterile eye-lubricating The high metabolic rate and high surface-to-volume ointment, especially in long-duration studies. ratio of mice means that they lose heat very quickly. • It is recommended to use a single injection while It is therefore imperative to avoid anesthetics such as delivering an injectable anesthesia to small barbiturates, which alter the animal’s ability to maintain rodents to reduce anxiety and ensure a stress-free core temperature (see PMIDs 18172330,15155266, induction and recovery. However, care must be 15027618). taken when mixing agents for a single injection Similarly, the animal should be warmed during operative to ensure safety and efficacy. procedures which open a body cavity and expose even greater surface area to ambient temperatures for heat loss. Body temperature should be monitored during heating to avoid increasing body temperature above 38ºC.

Effect of Core Temperature on Femoral Blood Flow in a 22 gram CD-1 Mouse: As the effect of progressive lower core temperatures in the respective flow traces demonstrates, temperature has a profound effect on femoral blood flow and must be monitored. Data, courtesy of M.F. Callahan, Dept. of Orthopaedic Surgery, Wake Forest University School of Medicine, Winston-Salem, NC

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Rodent Anesthesia Guidelines Cont.

Select References Alves HC, et. al. “Intraperitoneal propofol and propofol fentanyl, sufentanil and remifentanil combinations for mouse anaesthesia.” Lab Anim. 2007 Jul; 41(3): 329-36 (PMID 17640460) Blaudszun G, Morel DR. “Superiority of desflurane over sevoflurane and isoflurane in the presence of pressure-overload right ventricle hypertrophy in rats.” Anesthesiology. 2012 Nov; 117(5): 1051-61 (PMID 22929732) Gaertner, DJ, TM Hallman, FC Hankenson, MA Batchelder. 2008. Anesthesia and Analgesia in Rodents. Anesthesia and Analgesia in Laboratory Animals. Second Edition, Academic Press, CA. Gargiulo S, et. al. “Mice Anesthesia, Analgesia, and Care, Part I: Anesthetic Considerations in Preclinical Research.” ILAR J. 2012; 53(1): E55-69 (PMID 23382271) Hildebrandt IJ, et. al. “Anesthesia and other considerations for in vivo imaging of small animals.” ILAR J. 2008; 49(1): 17-26 (PMID 18172330) Janssen BJ, et. al. “Effects of anesthetics on systemic hemodynamics in mice.” Am J Physiol Heart Circ Physio. 2004 Oct; 287(4): H1618-24 (PMID 15155266) Roth DM, et. al. “ Impact of anesthesia on cardiac function during echocardiography in mice,” Am J Physiol Heart Circ Physiol 2002; 282: H2134-40 (PMID 12003821) Saha DC, et. al. “Comparison of cardiovascular effects of tiletamine-zolazepam, pentobarbital, and ketamine-xylazine in male rats.” J Am Assoc Lab Anim Sci. 2007 Mar; 46(2): 74-80 (PMID 17343357) Zeller A, et. al. “Mapping the contribution of β3-containing GABAA receptors to volatile and intravenous general anesthetic actions.” BMC Pharm 2007; 7(2) (PMID 17319964)

Ideal anesthetic agent Acknowledgement Thomas L. Smith, Ph.D., Bowman Gray School • Reliable of Medicine, Dept. of Orthopedic Surgery, Wake • Wide safety margin Forest University, Winston-Salem, NC. • Rapid onset/rapid recovery • Easy to administer & control • Nontoxic • Causes no physical impairment • Produces analgesia and muscle relaxation

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Aseptic Surgical Guidelines

The following protocol used at Bowman Gray School of Medicine is consistent with the Public Health Service guidelines. 1. Room Preparation 5. Draping the Animal a. The surgery table surface should be wiped with a. Drapes help to maintain a sterile field and a disinfectant before and after use. A bluepad/ preserve body temperature. drape should cover the table. 6. Closure of the Animal b. The use of a heating pad is recommended to prevent hypothermia and to aid in a quicker, a. Abdominal/thoracic body wall should be closed uneventful recovery. with absorbable suture material in a simple interrupted or similar interrupted pattern. 2. Animal Preparations b. Skin should be closed with non-absorbable, a. Remove hair from surgical site with clippers monofilament suture material in a simple and/or a depilatory. interrupted or similar interrupted pattern. b. The surgical site should be cleaned with an c. Sutures must be removed 7 to 10 days after antiseptic scrub followed by an antiseptic surgery to prevent inflammation and other solution (e.g. chlorhexidine or povidone iodine postoperative complications. scrub and solution, respectively). 7. Animal Recovery 3. Instrument Sterilization a. Recovery should occur in a warmed a. All instruments must be sterilized. The method environment. of choice will be determined by the surgical b. Animals should be observed closely until they instruments or devices being used. are able to maintain a sternal position and 4. Surgeon Preparation then every 6-8 hours until fully recovered. a. The surgeon and all others in the operating c. Post-operative antibiotics should be given after room must wear a surgical face mask prior surgery when justified by the investigator and to initiation of animal prep. The surgeon is the veterinary staff. required to wear sterile gloves; a cap and 8. Multiple Surgical Procedures sterile gown is also recommended. a. After the first surgery, the sterilized b. The surgeon must wear a scrub shirt. Non- instruments must be kept in a sterile tray surgeon personnel in the room must wear a lab containing 70-90% ethyl or isopropyl alcohol or coat or a gown over their street clothes. other acceptable solution. b. Sterile gloves must be changed between Aseptic Surgery surgeries. • Follow recommended guidelines & common sense. Healthier animals yield better science. Reference • Sterility: follow aseptic techniques; all instruments Principles of Proper Laboratory Use in Research and Teaching, Wake and gloves, mask, drapes, implants, sutures should Forest University, Bowman Gray School of Medicine, 1992 be sterile. • Use top quality surgical instruments: buying the instruments constitutes an initial investment, but the instruments will last for a long time if properly maintained.

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Surgical Recovery

Principles of Proper Laboratory Use in Research and Teaching, (Wake Forest Univ., Bowman Gray School of Medicine, 1992)

During the post-surgical period, a record must be kept in the room where the animal is housed. It should include a brief description of the surgical procedure, anesthetic used, time of induction, duration of surgical anesthesia, and time returned to cage. It should also include the findings of each physical examination during the recovery period. The post surgical medical record will be retained as a part of the animal’s permanent medical record. It is best to keep experimental notations in a separate location. For medical monitoring purposes, it is helpful to stage animals according to extent of recovery from surgery and anesthesia. The animal should be examined and the findings recorded according to the following schedule:

Stage 4 - Animal unconscious or semiconscious and unable to sit or maintain sternal recumbency. a. Examine and record findings no less frequently than every 2 hours. More frequent examination is recommended. Examples of notations include: i. Body temperature ii. Heart rate iii. Respiratory rate iv. Capillary refill time (record in seconds) v. Jaw tone (record resistance or no resistance) vi. Response to toe pinch (record withdrawal or no withdrawal) vii. Time of extubation b. Animal should be turned from side to side frequently to prevent dependent pulmonary congestion & edema. c. Ambient temperature should be adjusted (heat lamp or warming board) to bring body temperature to normal. Take care to not burn or over heat animal at this stage of recovery. The animal should be kept dry. d. The state of hydration should be assessed and fluids should be provided as necessary.

Stage 3 - Animal conscious & can maintain sternal recumbency to sit, but can not stand a. Examine and record findings so less frequently than every 6-10 hours depending on the nature of the surgery and the status of the animal. Examples of notations include: i. Body temperature until it becomes normal + 2°F. ii. Capillary refill time iii. Condition of the operative site b. Examine closely for other abnormalities c. Keep the animal dry and adjust the ambient temperature to bring the body temperature to normal d. Consider use of analgesic medication e. Professional judgement should be exercised in those cases in which there is difficulty in examinations every 6-10 hours. We recommend that one should be cautious in prolonging examinations of animals in stage 3.

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Surgical Recovery Cont.

Stage 2 - Animal can stand and move about but is not eating and drinking normally a. Examine daily and record findings. Examples of notations include: i. Body temperature ii. Hydration iii. Attitude (alert or depressed) iv. Activity (active or inactive) v. Food consumption vi. Water consumption vii. Condition of operative site b. Examine closely for other abnormalities c. Consider use of analgesic medication

Stage 1 - Animal active, alert, eating and drinking normally; skin sutures are in place a. Examine daily and keep a post surgical record of surgical site care until the sutures are removed. b. Sutures should be removed within 10-14 days of surgery.

Stage 0 - Animal normal and skin sutures removed. a. Specific post surgical care and record are no longer required.

Ideal Anesthetic Agent Acknowledgement Thomas L. Smith, Ph.D., Bowman • Reliable Gray School of Medicine, Dept. of • Wide safety margin Orthopedic Surgery, Wake Forest University, Winston-Salem, NC. • Rapid onset/rapid recovery • Easy to administer & control • Nontoxic • Causes no physical impairment • Produces analgesia and muscle relaxation

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PV Workbook RPV-3-wb Rev A 2014