Online-only appendix: Detailed technical information
I.
Catheterization
a. Choice of catheterization site: We catheterize the left femoral artery and vein
as opposed to the carotid and jugular. In our experience, mice catheterized via
the femoral site recover food intake more quickly. Disadvantages to the femoral
site include smaller blood vessels, and potential partial loss of use of the involved
leg.
b. Catheter preparation: MRE-025 tubing (Braintree Scientific, Braintree, MA) is
first cut to appropriate length (long enough to extend from femoral site, tunnel to
upper back, then rise from mouse to connection with swivel suspended above
cage). The tubing is heated in hot oil, slowly pulled to a narrow diameter, allowed
to cool, and cut at point where diameter is widest possible that will allow
placement in vessel. Tubing is then shaped into a J-form by threading into a
hollow metal J-shaped mold with an approximately 1cm diameter J curve,
submerging in hot water, followed by rapid cooling on ice. Formed catheters are
sterilized by ethylene oxide. Prior to surgery, arterial catheters are marked with
red ink to distinguish them from venous catheters after placement.
c. Surgery: Before surgery, catheters are filled with heparinized saline (20U/ml)
and weighed with standard 1cc syringes and doubled length of wire (792500; AM-Systems, Sequim, WA). Suture is prepared by separating, under a
microscope, strands of a 2-inch length of Ethicon 2-0 (3-0 metric) silk. Mice are
weighed, anesthetized using inhaled isoflurane, and transferred to a heated
platform under a dissection microscope. The upper back region is shaved using
antibacterial soap and scalpel, then cleaned using 70% ethanol. A small
horizontal incision through the skin is made posterior to ears, between scapulae.
The mouse is placed in supine position, immobilized using tape, and anterior
surface of left leg is shaved to lower abdomen and cleaned with 70% ethanol.
After vertical incision over femoral site, the artery and vein are dissected, with
visualization of operative field using micro-retractors made from paperclips taped
to platform. The operative field is kept moist using warm saline. The straight ends
of the catheters are tunneled to the upper back incision by threading through a
blunt 18G needle extending from the femoral site to the upper back site;
catheters are flushed with heparinized saline. The artery and vein are carefully
separated, and suture is drawn around each vessel individually at the proximal
end; suture ends are clamped off to the side. Using short suture, the distal artery
and any larger branches in operative field are tied off. Using the proximal suture,
tension is applied to artery until blood flow is impeded. A small incision is made in
the distal artery, the beveled arterial catheter is inserted and tied in place with
suture, without closing off the catheter. Tension is released from clamped suture.
Catheterization of the femoral vein occurs by the same method except the
catheter is not beveled. The operative site is coated with superglue to fix the
catheters into position relative to surrounding structures. Buprenorphine 0.1
mg/kc SQ and penicillin G 12000U IM are administered while glue hardens. The
wound is closed using 6-0 silk suture, using approximately 7 separate sutures,
and cleaned with iodophore. Mouse is returned to prone position. Catheters are
checked by syringe withdrawal and flush. Looped wire is tied very tightly to the
muscles of the upper back and neck, near catheter exit site, to stabilize and
protect catheters and provide torque to turn swivel (see below). Catheters are
tied to wire, and catheters are checked by syringe withdrawal and flush.
Catheters are taped tightly to the wire at two inch intervals, using approximately
four pieces of tape, starting immediately above neck. The neck incision is
superglued and closed.
d. Recovery: After recovery from anesthesia, mouse is transferred to caging unit
(see below), wire and catheters are attached to swivel (see below) and mouse is
monitored for blood pressure, heart rate, movement, and nest formation.
II. Tether system
a. Choice of tether system: In our experience, mice bite tether jackets, and their
activity is hindered by commercial spring tethers, so we use simple thin wire
stabilization instead. Mice are housed in round-bottom cage units (Instech,
Plymouth Meeting, PA) and are given enough slack that they can use the entire
cage bottom, so in fact “tether” is somewhat a misnomer, as they have full
freedom of movement. The purpose of the wire is to impart enough stiffness to
allow the mouse to turn the swivel. Without the wire, the mouse turns freely, the
catheters remain fixed to the swivel which doesn’t turn, and the catheters
become knotted.
b. Swivel: We use an ultra low-friction 360° dual channel stainless steel swivel
designed for mice (375/D/22QM; Instech, Plymouth Meeting, PA). These swivels
can be used repeatedly; some of ours have been in use continuously for more
than a year. We suspend the swivel above the cage using a simple arrangement
of square-bottom laboratory stands and clamps. The swivel ports are connected
to PE-50 tubing (Becton Dickinson #427411); short lengths, at bottom ports, are
connected to 26G needles for attachment to mouse catheters, and longer
lengths, at top ports, are connected to continuous flush syringes (see below).
III. Housing and husbandry
a. Light/dark: Mice are kept in light- and sound- proof boxes (BRS-LVE
Environmental cubicle) with a small internal light source kept on a 12 hour
light/dark cycle. Syringe pumps are kept outside the cubicle to maximize space
and reduce noise and light.
b. Food, water: Mice have ad libitum access to food and water at all times. Food
pellets are weighed daily to quantify chow caloric intake.
c. Caging units: Mice are kept in small animal round cage enclosures (S-Tank,
Instech, Plymouth Meeting, PA) with sawdust bedding and a nestlet. Plumping of
the nestlet has proved to be an extremely valuable measure of animal
postoperative health.
d. Monitoring: Mice are checked a minimum of twice daily for general health (blood
pressure, heart rate, nestlet plumping, movement, signs of bleeding). In general,
our postoperative recovery rate is greater than 90%.
IV. Catheter maintenance
a. Hemodynamic monitoring: The arterial catheter is connected to the vertical
channel of the swivel. The distal port of the swivel is attached to PE-50 tubing
connected via a 22G needle to a pressure transducer (CDExpress, Owens and
Minor, Greensburg PA) connected to a 1cc syringe continuously flushing
heparinized saline (20U/ml) at a rate of 7µl/hr. The pressure transducer cable is
connected to a recorder and amplifier system (Dataq Instruments, Akron, OH).
The digital output is continuously displayed on a computer monitor using Windaq
software (Dataq Instruments). This information is extremely helpful in monitoring
mice and catheters. The mean blood pressure and heart rate give information
regarding the health of the animals, and the tracing amplitude indicates the
degree of patency of the arterial catheter.
b. Continuous flush: Both arterial and venous catheters are connected, postswivel, to 1cc syringes containing heparinized saline (20U/ml) which flush
continuously at a rate of 7µl/hr, driven by a syringe pump with a multi-syringe
adaptor (R99-EM with R-ACC-6, Razel Scientific Instruments, St. Albans, NY).
This continuous flush is extremely important for maintaining catheter patency.
c. Manual flush: If the arterial pressure tracing shows reduced amplitude, this may
indicate that the arterial catheter has become clogged. If caught early, manually
flushing the arterial catheter using a 1cc syringe with 26G needle can allow reopening of the catheter. Care must be taken to avoid infusing air into the mouse.
V. Arterial blood sampling
a. Obtaining sample: To obtain blood for measurement of blood glucose and
plasma for insulin or corticosterone assays, we use the arterial catheter. Using a
small hemostat clamp, the arterial line is clamped near the swivel. The catheter is
removed from the swivel tubing, accessed using a 1cc syringe with 26G needle
containing heparinized saline, and blood mixed with saline is removed from the
catheter. After an appropriate volume is removed, the catheter is reclamped, a
clean insulin syringe is attached, approximately 100µl whole blood is removed,
the catheter is reclamped, the original syringe is reattached, the dead volume
blood and saline is reinfused, and the catheter is reattached to the swivel.
b. Returning red blood cells: After measuring blood glucose using a glucometer
(Ascencia Elite XL, Bayer, Tarrytown, NY), the remainder of the blood is
centrifuged in an eppendorf tube, and the plasma is transferred to a clean tube
and stored at -80 degrees. The red blood cell pellet is resuspended using sterile
heparinized saline (20µl of 100U/ml) and reinfused into the mouse to prevent
anemia from frequent sampling.
VI. Infusions
a. Setup: Sterile saline or glucose solutions, mixed with sterile bromodeoxyuridine
(500µg/ml), are transferred to a 10cc syringe. The syringe is connected via a 22G
needle to a length of PE-50 tubing. After priming the tubing with solution, the
syringe is driven by a syringe pump (BS-8000, Braintree Scientific, Braintree,
MA). The PE-50 is connected to the side port of the swivel. 35µl is delivered as a
bolus, to account for the dead space between swivel and mouse. All tubing is
checked carefully for leaks and air bubbles.
b. Infusion: The infusion pump is checked twice daily to make sure it is powered on
and pumping. The volume infused is recorded daily.
VII. Troubleshooting and potential technical problems
a. Surgery: An experienced surgeon is required to catheterize mice. Optimal
catheterization requires that maximally sized catheters are placed, with a
minimum of bleeding. The duration of the operation is also important to the
outcome; we find that surgeries lasting more than 60-90 minutes can impact on
the speed of recovery and health of the mouse.
b. Tether: if the wire is not tightly attached to the neck muscles, or if the wire
becomes detached from the swivel, the catheters become tangled. Our mice are
quite active, even in the first postoperative day, and can tangle catheters quickly
if the swivel is not engaged. Also, if any objects or catheters interfere with the
free turning of the swivel the catheters will become knotted. The most frequently
encountered problem is with infusion catheters draping down into the path of the
turning mouse catheter/wire assembly. This is easily solved by routing the
infusion catheter above and behind the swivel clamp, or taping it to the ceiling of
the box.
c. Catheter maintenance: Keeping arterial catheters patent is fairly labor intensive.
We check the catheters at least twice daily by monitoring the blood pressure
tracing amplitude and/or manual withdrawal and flush. Any leaks in the arterial
system can result in clogging of the catheter, so each time the mouse is checked
the entire system is observed for leaks while flushing at a higher rate. The most
common leak sites are where catheters connect to needles, including at the flush
syringe, pressure transducer, and swivel. If a leak is found, the punctured
catheter is clamped, cut, and reattached.