Free-Hand Intracerebroventricular Injections in Mice

Summary

Here, a simple and rapid approach for performing intracerebroventricular injections in mice using a free-hand approach (that is, without a stereotaxic device) is described.

Abstract

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. An intracerebroventricular (ICV) injection allows the widespread delivery of the experimental agent throughout the brain (particularly in the structures near the ventricles). Here, methods for making free-hand ICV injections in adult mice are described. By using visual and tactile landmarks on the heads of mice, injections into the lateral ventricles can be made rapidly and reliably. The injections are made with a glass syringe held in the experimenter's hand and placed at approximate distances from the landmarks. Thus, this technique does not require a stereotaxic frame. Furthermore, this technique requires only brief isoflurane anesthesia, which permits the subsequent assessment of mouse behavior and/or physiology in awake, freely behaving mice. Free-hand ICV injection is a powerful tool for the efficient delivery of experimental agents into the brains of living mice and can be combined with other techniques such as frequent blood sampling, neural circuit manipulation, or in vivo recording to investigate neuroendocrine processes.

Introduction

The delivery of experimental agents, such as drugs¹, viruses², or cells³, to the brain is often necessary for neuroendocrine research. If the agent does not readily cross the blood-brain barrier or the experimental objective is to specifically test the central effects of the agent, it is important to have a reliable method for delivering injections into the brain. Moreover, injection into the intracerebroventricular (ICV) space provides the opportunity to distribute the agent widely in the brain and provides a large target area, thus increasing the likelihood of successful injection².

A common method for making ICV injections involves the placement of a permanent indwelling cannula. In this approach, a stereotaxic frame is necessary to position the commercially available or custom-made cannula, as the cannula is glued or cemented in place. Often, upon recovery, a supraphysiological dose of angiotensin II is administered through the cannula, and if drinking behavior is immediately observed, then the cannula is considered correctly placed⁴. This approach has many advantages, including the ability to perform long-term infusion and the ability to inject the same animal multiple times; additionally, if angiotensin II is employed, correct placement can be confirmed prior to the administration of experimental compounds. However, there are some limitations to placing a permanent cannula, including the requirement for expensive equipment (stereotaxic frame), the possibility of damage to the cannula after placement (e.g., mice can chew on the cannula of a cage mate), and the possibility of infections around the permanent cannula. Single ICV injections can be made with the use of a stereotaxic frame³, which, although effective, requires substantial exposure to anesthesia and, thus, may obscure some acute physiological and behavioral effects of the treatment. Additionally, the placement of mice in a stereotaxic frame requires substantial training to achieve stable placement and prevent the rupturing of the ear canals.

Here, an established method for making free-hand injections in mice is described. This method is based on previous reports^5,6. The advantages of this technique are that it is simple, rapid, and does not require specialized equipment such as a stereotaxic frame. As described below, this procedure involves manipulating a glass syringe relative to landmarks on the mouse head to make the injections, which can be done rapidly and, thus, requires only a few minutes of gas anesthesia on the experimental day.

Protocol

All the procedures were approved by the Colorado State University (#3960) and University of California San Diego Institutional Animal Care and Use Committees, where the representative data were collected (S13235, PI Kellie Breen Church). Data from five adult female and two adult male C57/BL6 mice (9-16 weeks old) are depicted in the representative data section. Female mice were ovariectomized 3-4 weeks prior to ICV injection and blood collection as described previously⁷. Prior to experimentation, these mice were housed with a 12 h light/12 h dark light cycle and had free access to feed and water in accordance with the Guide for the Care and Use of Laboratory Animals.

1. Performing craniotomy

NOTE: Craniotomy can be performed one or more days prior to the actual injection, which makes the injection process quicker on the day of the experiment.

1. Prepare the materials for the craniotomy as described below.
   1. Place a clean bench pad or drape material on the work surface. Tape the nose cone of an isoflurane vaporizer to the work surface (close to, but facing away from, the experimenter; see Figure 1A).
   2. Place silastic tubing on the end of a sterile sharp 18 G needle so that ~1 mm of the needle tip protrudes. Place silastic tubing on the end of a sterile blunt needle so that ~1 mm of the needle tip protrudes.
      NOTE: Separate and sterile pre-packaged needles (sharp and blunt) should be used for each mouse.
   3. Gather an electric shaver, iodine scrub, sterile gauze, and alcohol pads for preparing the injection site.
2. Practice moving a needle 2 mm lateral and 1 mm caudal as described below.
   1. Use a ruler and a pen to mark a sheet of paper with a starting point (this will be bregma), a mark 2 mm to either the left or right, and another mark 1 mm above the last one (this will be the injection site; see Figure 1B).
   2. Practice moving the needle from the starting point to mark point 1 to mark point 2 (bregma to injection site) until confident that the movement is repeatable without guides.
      NOTE: The coordinates described here are effective in adult C57/BL6 mice, but other strains or age groups may require adjustment.
3. Prepare the mouse for craniotomy as described below.
   1. Place the mouse in an induction chamber and anesthetize the mouse with 3%-4% isoflurane in medical-grade oxygen. With practice and familiarity with the procedure, reduce the total duration of isoflurane exposure to less than 10 min for the craniotomy procedure.
      CAUTION: Isoflurane exposure is hazardous to human health; use an approved and inspected isoflurane vaporizer in a well-ventilated area with means of scavenging waste gases.
   2. Once the toe reflex is absent, shave the head of the animal. Apply eye lubricant.
   3. Place the mouse on the work surface with the head in the nose cone to maintain anesthesia.
   4. Administer an analgesic agent, such as buprenorphine (0.6-0.8 mg/kg mouse body weight, subcutaneously), as directed by the supplier.
   5. Clean the injection site by wiping the head with sterile gauze dipped in an iodine solution (perform three scrubs), and then wipe with an alcohol scrub pad (perform three times).
      ​NOTE: Remove the fur and clean the skin around the injection site and use sterile instruments and needles, as recommended by the American Veterinary Medical Association to prevent infections.
4. Firmly hold the head of the mouse with the non-dominant hand. Position the head as flat on the work surface as possible.
5. Locate the site of the injection.
   1. Use the prepared sharp 18 G needle to first identify bregma; for this, drag the needle across the skin of the head along the midline, moving in the rostral-caudal plane. Imagine an equilateral triangle in which the two vertices are the eyes and the third vertex is the approximate location of bregma (see Figure 1C).
   2. From bregma, move the needle 2 mm lateral and 1 mm caudal to the site of injection.
6. While holding the needle vertically, firmly push the needle through the skin and bone until the tubing is flush with the skin.
7. Retract the needle, rotate the needle, and press through the skin and bone again. Repeat the process until a small hole is created in the bone.
8. Use the blunt needle to check that a sufficient hole has been produced in the bone. Aim to make an opening in the bone large enough for the blunt needle to pass through.
9. Remove the mouse from the nose cone, clean off any blood from the injection site with sterile gauze, and place the mouse in a cage on a cage warmer until awake. Since the opening made by the craniotomy is small (18 G needle), the site does not need to be closed with a suture or wound clips.
   NOTE: The opening produced here is small (18 G), so many institutions consider this procedure an injection as opposed to a surgery. Moreover, although an opening through skin and bone to the brain is made, holding the skin taut results in the holes not aligning, which likely aids in preventing skin flora or cage bedding from contacting the brain.

2. Making the injection

1. Prepare the materials for the injection as described below.
   1. Place a clean bench pad or drape material on the work surface. Tape the nose cone of an isoflurane vaporizer to the work surface (close to, but facing away from, the experimenter; see Figure 1A).
   2. Attach a 10 mm long 27 G needle with a 45° bevel onto a 5 µL glass syringe. Place silastic tubing on the needle so that 3.5 mm of the needle tip protrudes; use laboratory tape to hold the tubing to the syringe body (see Figure 1D).
   3. Prepare the injection media (drug, virus, or other liquid for injection) in a tube. For the representative results in this study, sterile isotonic saline was injected.
      NOTE: Select a tube that permits access by the glass syringe and needle, such as a 2 mL microcentrifuge tube.
   4. Draw 3 µL of injection media into the glass syringe. First, draw more than the desired volume, and eject until 3 µL remains in the syringe.
   5. Gather iodine scrub, sterile gauze, and alcohol pads for preparing the injection site.
   6. Start a laboratory timer in count-up mode and place the timer so that it is visible to the experimenter while making injections.
   7. Practice moving a needle 2 mm lateral and 1 mm caudal as performed on the previous day.
2. Prepare the mouse for injection as described below.
   1. Place the mouse in the induction chamber and anesthetize the mouse with isoflurane. Once the toe reflex is absent, apply eye lubricant, and place the mouse on the work surface with the head in the nose cone.
   2. Clean the injection site with three wipes each of iodine and alcohol. Identify the injection site with an 18 G needle (or blunted needle) as described in step 1.8. The hole created during craniotomy should be detectable.
      NOTE: If the craniotomy has not been performed in advance, or if the hole in the skull is not detectable, the craniotomy can be performed here.
3. Perform the injection with a glass syringe as described below.
   1. Firmly hold the head of the mouse with the non-dominant hand. Position the head as flat on the work surface as possible.
   2. Place the needle of the glass syringe through the hole in the skull until the tubing placed over the needle prepared in step 2.1.2 rests on the skin of the mouse.
   3. Hold the syringe as vertically as possible, paying attention to both the coronal and sagittal planes. Slowly inject the media over a period of 1 min.
   4. Hold the syringe and needle in place for another minute after the completion of the injection to minimize backflow. Slowly retract the needle from the head of the mouse.
   5. Remove the mouse from the nose cone, clean off any blood from the injection site with sterile gauze (if there is any present), and place the mouse in a cage on a cage warmer until awake.

3. Confirming the injection location

1. Deeply anesthetize and perfuse the mouse with saline and 4% paraformaldehyde in phosphate-buffered saline (PBS) as previously described⁸. Store the brain in 30% sucrose in PBS at 4 °C until the brain sinks (typically 2 days).
2. Cut 20-50 µm coronal sections on a cryostat as described previously⁹. While sectioning, observe the injection tract in the tissue block. Record whether the injection tract clearly intersects the lateral ventricle.

Results

When performed successfully, this technique allows the rapid delivery of an experimental agent into the ventricular system. A luteinizing hormone (LH) pulse profile from an ovariectomized mouse that received an ICV injection of 3 µL of sterile isotonic saline, the vehicle for many pharmacological compounds, is shown in Figure 2A. This example demonstrates that brief exposure to gas anesthesia and the injection of 3 µL of fluid into the ventricular system alone did not alter the pul...

Discussion

Here, a simple and effective means for making ICV injections in mice is described. Since this technique does not require a stereotaxic frame, this approach for the central delivery of drugs and experimental agents is accessible to more researchers. Additionally, this approach is relatively high throughput since the preparation and injection procedure can be performed quickly.

Since this procedure requires the manipulation of needles and a glass syringe by hand using approximate distances, it i...

Materials

Name	Company	Catalog Number	Comments
18-gauge blunt needles	SAI Infusion	B18-150
18-gauge needles	BD Medical	305195
Alcohol pads	Fisher Scientific	22-363-750
Bench pad	Fisher Scientific	14-206-62AC22
Betadine solution	Fisher Scientific	NC1696484
Buprenorphine	Patterson Vet Supply	07-892-5235	Controlled substance
Eyelube	Fisher Scientific	50-218-8442
Glass syringe	Hamilton	7634-01
Injection needle	Hamilton	7803-01	27 gauge, Small Hub RN needle, point style: 4, Needle length: 10cm, Angle: 45
Isoflurane	Patterson Vet Supply	07-893-8441
Isoflurane vaporizer	Vet Equip	V-10
Laboratory Tape	VWR	89098-128
Medical grade oxygen	Airgas	OX USPEA
Paraformaldehyde	Millipore-Sigma	8.18715.1000
Phosphate Buffered Saline	Fisher Scientific	J67802.K2
PulsaR Software	Open source, University of Otago		See ref 9
Ruler	Fisher Scientific	12-00-152
Silastic tubing (0.040" I.D.)	DOW	508-005
Silastic tubing (0.078" I.D.)	DOW	508-009
Sterile saline	VWR	101320-574
Sucrose	Fisher Scientific	S5-500