TITLE: Structural MRI Using the University of Washington 14T Vertical Bore Bruker MRI
AUTHORS: Yoni Browning, Galen Lynch, Josh Siegle

[DESCRIPTION]
This protocol describes the process of obtaining pre-insertion MRI images using the University of Washington 14T vertical bore Bruker MRI. These structural scans allow for precisely targeting acute electrophysiology recordings in head-fixed mice beyond what is possible using skull landmarks alone.

[BEFORE_START]
Complete the whole hemisphere craniotomy surgery, but substitute a zirconia headframe for the standard titanium version.

Titanium headframes will cause MRI artifacts. This procedure requires a zirconia headframe. Prep the zirconia headframe for surgery by coating the outside of each of the fiducial holes with a thin layer of dental wax. This will prevent Metabond from wicking into the headframe holes during implantation, which would limit our ability to register the MR scan. (See "Prepare for MRI scans" for details about the headframe).

After surgery, recover the mouse according to the surgical and IACUC protocols.

[GUIDELINES]
Only perform this procedure in accordance with IACUC and veterinary requirements.

[STEPS]
SECTION: Prepare MnCl2 solution for injection.
1. Mix 100 millimolar (mM) MnCl2 solution. (These steps are from section 3.1.1 of Massaad and Pautler (2011).)
 

SECTION: Prepare MnCl2 solution for injection.
1.1. Create 100 mL of a 100 millimolar (mM) bicine buffer solution by mixing 1.63 g bicine into 100 mL MilliQ water in a beaker.

SECTION: Prepare MnCl2 solution for injection.
1.2. Place clean stir bar into the bicine solution and place on a stir plate. With a calibrated pH meter, bring the bicine buffer solution to pH 7.4 using 1 Molarity (M)  NaOH.

SECTION: Prepare MnCl2 solution for injection.
1.3. Sterilize the bicine buffer solution using a   syringe filter into a sterile container that can hold at least 100 mL.

SECTION: Prepare MnCl2 solution for injection.
1.4. In a hood, aliquot 5 mL  of the sterile bicine into a sterile container. Using sterile tools, measure 98.95 mg of MnCl2 with a microbalance. Mix the MnCl2 into the bicine aliquot.

SECTION: Prepare MnCl2 solution for injection.
1.5. Using a sterile syringe and needle, transfer MnCl2 solution to sterile septum vial using sterile techniques.
SECTION: Prepare MnCl2 solution for injection.
2. The injectable should be refrigerated at 4 °C. Discard when expired according to local IACUC. Label accordingly.

SECTION: Inject mice with MnCl2 solution, 18-24 hours prior to imaging
3. Weigh mouse.
SECTION: Inject mice with MnCl2 solution, 18-24 hours prior to imaging
5. Using sterile technique, draw the volume of MnCl2 injectable calculated above into an insulin syringe. Perform an intraperitoneal (IP) injection of the MnCl2 solution.
 

SECTION: Inject mice with MnCl2 solution, 18-24 hours prior to imaging
6. It is normal for mice to be lethargic/uncomfortable after MnCl2 injection. These symptoms typically resolve themselves within a few hours. Work with your IACUC and veterinary staff to identify symptoms of concern and establish protocols to aid in recovery. We recommend the following:
Provide thermal support to the mouse for a minimum of 30 minutes using an electric heating pad or chemical heat pack.
Monitor mice for signs of unusual pain/discomfort, and to ensure continuous improvement.
For mice not on water restriction, provide fluid support to the mouse via subcutaneous injection of 0.25-.5 mL of sterile LRS (Lactated Ringers Solution) or saline.


SECTION: Prepare for MRI scans
7. Place each mouse in a disposable transport cage. Transport the mice to the imaging facility. For transport, transfer cages should be placed in a large plastic bin that is secured with ratchet straps in the back of an approved cargo van.
SECTION: Prepare for MRI scans
8. Confirm the mouse ID number by matching ear tags/tattoos to the ID number on the cage.
SECTION: Prepare for MRI scans
9. Anesthetize the mouse with 2-5% isoflurane using an induction chamber.
SECTION: Prepare for MRI scans
12. Fill the fiducial holes in the mouse’s headframe with degassed white petroleum-based eye lubricant (e.g. Systane Nighttime lubricant) or Vaseline using a 25 gauge blunt-tipped syringe. To do this, you will likely need to remove any protective wax placed over the holes prior to surgery; this can be done using any sharp-tipped needle. Backfill the holes, taking care to avoid bubbles in the holes as these will limit the ability to see the holes in the final scan. Good technique for doing this is to perturb the Vaseline mixture as you fill by moving the syringe in the hole so that any bubbles that do form are forced out. Additionally, spread eye lubricant or Vaseline onto the lower plane of the shank of the head bar.

During post-hoc analysis, these fiducial fills will be used to align the scan to the headframe holder on the rig. The lower plane of the headframe and other geometry can be used to augment incomplete hole fills. See Registering MRI Volumes to an In Vivo Electrophysiology Rig for more information. If you are having trouble localizing the headframe in your scans, it can be useful to spread eye lubricant around other salient geometric features on the headframe so that you have more data for alignment.

 

SECTION: Prepare for MRI scans
10. Apply eye lubricant to the mouse’s eyes.
SECTION: Perform the primary MRI scan
15. Perform the full 3d scan. For the UW MRI scanner, typical parameters are as follows. Work with your scanner operators to validate that these parameters are correct for your equipment and needs. Your MRI Technician will set these parameters and can load parameters from previous scans.

Parameters:
Scan Type: RARE (fast spin echo)
Resolution: 100 µm isotropic
Number of Averages: 2
RARE Factor: 4
Echo Time: 5.333 ms
Effective TE (time to echo): 10.666 ms
Repetition Time: 500 ms
 

SECTION: Perform the primary MRI scan
14. The trained MRI technician will operate the scanner. Work with them to achieve the following. Note that the order of these steps may change on different hardware/software, that some steps may be automated, or that additional steps may be required depending on the equipment used. If any steps here are unclear, consult the manual for your equipment.
Adjust the position of the mouse in the scanner to achieve the best scan quality. Ideally, the brain will be as centered in the scan as possible while still allowing the headframe holes to be visualized.
Perform matching and tuning (consult your MR technician and/or your scanner's manual for details on this step).
Determine the scan volume desired, being sure to include both the brain and the positioning holes in the headframe. To do this, we typically collect axial and coronal scans with large slice thickness and use these to confirm that all needed features are included.
The readout direction for the high-resolution 3D scan should be chosen to align with the mouse's AP axis to avoid signal aliasing. In the non-readout axes, place saturation bands around the scan volume as needed to limit aliasing. This will allow for a smaller scan volume while minimizing aliasing. 
We recommend taking a low-resolution scan prior to the "full" 3D scan list in the next step. This will allow you to confirm that the headframe holes have adequate fill and that the scan volume is correct.
If the hole fills are incomplete or show artifacts, take the mouse out of the scanner and repeat  , being sure to avoid air bubbles in the Vaseline.

SECTION: Perform the primary MRI scan
16. Remove mouse from scanner (done by MRI operator) and recover mouse with thermal support. Return the mouse to the transport cage once the mouse begins to ambulate.
SECTION: Perform the primary MRI scan
17. Repeat scanning procedure for all mice in a group, sanitizing any surfaces the mouse contacts after each scan.
SECTION: Perform the primary MRI scan
18. Transfer the data to a USB drive.
SECTION: Transport mice back to home vivarium
20. Transfer data to appropriate storage location.
SECTION: Prepare for MRI scans
13. Load the mouse into the RF coil, securing the headframe to a holder if using one. 


 


SECTION: Transport mice back to home vivarium
19. Transport mice back to home institute and follow IACUC protocols for returning mice.
SECTION: Inject mice with MnCl2 solution, 18-24 hours prior to imaging
4. Calculate the dose of MnCl2 injectable needed for 66 mg/kg of manganese. Multiply the mouse weight by 3.3348 µL. If the mouse has an implanted headframe, subtract the weight of the headframe from the mouse weight before calculating the dose.
SECTION: Prepare for MRI scans
11. Secure the MRI-compatible nose cone. While some scanners have prefabricated cones, the headframe on our mice interferes with the commercially available options from Bruker. We find that a rubber pipe bulb, cut to accommodate the mouse's nose and teeth, works well. This can be secured to the mouse's face using strips cut from thin surgical tubing. The image below illustrates this setup and how it is attached to the mouse as well as to the anesthesia system.



 


SECTION: Prepare for MRI scans
13.1. Our headframe holder for the Bruker 14T Vertical bore magnet is illustrated below, though modifications may be needed to affix other headframes or other scanners. In our case, we use a plastic nut (A) and screw (B) to affix a plastic holding bar (C) to a stabilization bolt hole on the mouse's headframe (D). The plastic bar is positioned roughly in line with the protruding handle on the mouse's headframe. The screw should be tight enough that the mouse does not move, but care should be taken not apply to much force as the plastic interface can strip easily. The exact position here is not strictly important - fine alignment will use the fiducial holes filled above; this setup is just to prevent the mouse from moving during the scan.

 

SECTION: Prepare for MRI scans
13.2. To position the animal in the center of the coil, it can be helpful to measure the distance from the center of the animal's brain to a bolt on the headframe holder. Because these bolts will stick out of the coil once the animal is loaded, they can be used to approximately center the mouse's brain in the coil.

 

SECTION: Prepare for MRI scans
13.3. Because the headframe is wider than the mouse's head, the animal needs to be loaded into the RF coil at an angle. The mouse should be tilted so that the holder is pointing straight, and the animal's head is tilted to the right. The gas line for the anesthesia should be passed through the coil ahead of the mouse and should sit under the animal when it is loaded into the scanner (if it is placed to the side, it can cause "fold over" or "wrap-around" imaging artifacts in the scan. Take care when loading and unloading the animal into the coil - rough handling here can put unnecessary strain on the animal's neck. 

 

SECTION: Prepare for MRI scans
13.4. Once the animal is fully loaded into the coil, a trained MRI technician can finish securing the animal and place a breathing pillow to monitor respiration during the scan.