Home‎ > ‎Protocols‎ > ‎

M Aref and J Handbury DCE MRI Protocol


The authors are not responsible for anything that happens to anyone or anything given the use of this protocol. This is merely a guide based on our successful experiments, no more no less. A single person can do this entire experiment but it is a lot easier to do with two sets of hands. NOTE: Gloves should be worn at all times when handling animals. Use an alcohol wipe to clean all rubber stoppers before withdrawing volumes.


The anesthesia used is a solution of 68 mg ketamine plus 14 mg xylazine plus 2.3 mg acepromazine per kg body weight. Use a long larger bore needle to withdraw the appropriate amount of anesthetic. Use a short small bore needle to inject. There are two techniques for inducing animal anesthesia:

  1. If the researcher is a novice with animal handling, a half capful of Halothane can be poured on Kim wipes, placed lightly crumpled, in the base of a large bell jar. A grate or other fenestrated surface should be placed over the Kim wipes to protect the animal. The animal should be introduced and the bell jar closed. NOTE: At all times the halothane bottle should remain in a hood. If you smell a sickly sweet solvent odor you are breathing in Halothane (which is hepatotoxic and teratogenic). The animal should be immediately be removed and injected when it does not respond to the bell jar being shaken.
  1. For lower intra-experimental fatality rates, do not use the above method. First, spread a disposable drape followed by a cloth towel. Then remove the animal from its cage and place it on the towel. Wrap the towel firmly but gently around the animal and hold in place, leaving the hindquarters free. Make sure the head is securely covered. Using the non-dominant hand to restrain the animal, inject with the dominant hand (see picture to right). Return the animal to its cage to allow the anesthetic to take effect. In either case an intramuscular (IM) injection of anesthetic solution is used. For best results inject half the dose in each thigh, laterally. Once the animal is non-responsive, cover eyes liberally with Duratears eye petroleum jelly and be sure to keep animal warm.

IV Access

Using the paw and tail pinch test insure the animal is non-responsive. Place the rat's tail in a beaker of warm (it should feel comfortable to your finger) water for two (2) minutes (see picture to left). Immerse the entire tail. Prepare a disposable drape and IV supplies. The catheter used is a 25G × 3/4" (0.50 × 19 mm) with 12" (30 cm) of tubing. A long, larger bore needle should be used to draw up 0.5 mL of 2 units per 100 mL of heparinized saline and the catheter should be filled. Two separate long, larger bore needles should be used to draw up (a) 0.5 mL of 2 units per 100 mL of heparinized saline and (b) 0.3 mmol/kg of 0.5 mmol/mL GdDTPA (Magnevist, Berlex Imaging) (http://www.berleximaging.com/magnevist/purchasing_info.html), respectively. Remove the rat's tail from the beaker and rapidly but gently dry. Quickly clean tail with alcohol wipe and place rat on its side. The tail veins run laterally down either side of the rats tail. Following vasodilation due to the water bath the tail veins should appear as faint dark blue lines approximately 0.5 mm in width. Starting at point approximately 3 cm from the most distal part of the tail (i.e. the tip), try to insert the needle into the tail vein. The catheter should be bevel up and should be inserted at a shallow angle. Flatten the angle so that there is maximal venous luminal area. Do not go perfectly horizontal. There are three signs to a successful venous puncture:

  1. There should be a slight "pop" as the needle pierces the vein
  1. A visible "flash" of blood should appear in the tube. If the BP is too low for a spontaneous "flash", one can be elicited by drawing gently back on the syringe. If there is no blood in the catheter tubing then the needle is NOT in the vein. Securely tape the catheter in place using surgical tape. Place one piece of tape on the opposite side of the tail from the catheter with the sticky side toward the "butterfly wings". Wrap securely but gently around the catheter. Take a second piece of tape and apply it perpendicularly to the first covering the needle and injection site. Now,
  1. Remove syringe with heparinized saline. Attach syringe with 3 mL of normal saline. Inject approximately 0.5 mL. Injection should be smooth and no pallor or edema should occur around injection site.
  1. Now attach the syringe with the contrast agent to the catheter, if the volume is less than 0.4 mL, inject slowly into catheter tubing. Waste the remainder of the normal saline in the sink except for a flush volume of 0.5 mL. Attach normal saline syringe to catheter but do not inject.


The animal and a 1 mM Gd-DTPA reference are loaded in the bed. The tail and IV with attached flush syringe are carefully draped along the length of the bed. The bed is loaded in the coil. Two microwavable heating pad (www.rattusetal.com) are placed in the bore, one in front of and one behind the coil.

T1-weighted GEMS

TR = 63 ms

TE = 4.3 ms

Flip angle = 800 (1000 microseconds 44 dB Gaussian)

FOV = RO 24 cm / 512 × PE 6 cm / 128

Averages = 2

No. of Slices = 7

Slice thickness = 2 mm

Orientation = coronal

BW = 105 Hz

The above image sequence is repeated for approximately 30 minutes (112 data sets). The first data set is the preinjection data, contrast agent injection occurs at the beginning of the second data set. That is, the 0.5 mL flush volume is pushed at this time.