Elimination of Serotonergic Neurons by Stereotaxic Injection of 5,7-Dihydroxytryptamine in the Dorsal Raphe Nuclei of Mice

Summary

This protocol describes a dorsal raphe nucleus (DRN)-lesioned mouse model (>90% survival rate in experimental mice) with stable loss of dorsal raphe serotonergic neurons by stereotaxic injection of 5,7-dihydroxytryptamine into the DRN using an angled approach to prevent injury to the superior sagittal sinus.

Abstract

Stereotaxic injection has been widely used for direct delivery of compounds or viruses to targeted brain areas in rodents. Direct targeting of serotonergic neurons in the dorsal raphe nucleus (DRN) can cause excessive bleeding and animal death, due to its location below the superior sagittal sinus (SSS). This protocol describes the generation of a DRN serotonergic neuron-lesioned mouse model (>90% survival rate) with stable loss of >70% 5-HT-positive cells in the DRN. The lesion is induced by stereotaxic injection of a selective serotonergic neurotoxin 5,7-dihydroxytryptamine (5,7-DHT) into the DRN using an angled approach (30° in the anterior/posterior direction) to avoid injury to the SSS. DRN serotonergic neuron-lesioned mice display anxiety-associated behavior alterations, which helps to confirm success of the DRN lesion surgery. This method is used here for DRN lesions, but it can also be used for other stereotaxic injections that require angular injections to avoid midline structures. This DRN serotonergic neuron-lesioned mouse model provides a valuable tool for understanding the role of serotonergic neurons in the pathogenesis of psychiatric disorders, such as generalized anxiety disorder and major depressive disorder.

Introduction

Serotonin, or 5-hydroxytryptamine (5-HT), is an important neurotransmitter mainly produced in the intestines and brain and impacts a variety of psychological functions. In the central nervous system (CNS), the serotonergic system plays a central role in the regulation of mood and social behavior, sleep and waking, appetite, memory, and sexual desire. In the CNS, serotonin is synthesized by serotonergic neurons, which can be separated into the following two groups: the rostral group, which has ascending projections innervating virtually the whole brain; and the caudal group, which mainly projects to the spinal cord¹. The rostral group, which contains about 85% of serotonergic neurons in the brain, is composed of the caudal linear nucleus, median raphe nucleus, and DRN, in which the largest population of serotonergic neurons in the brain is located.

Dysregulation of the serotonergic system is generally believed to be linked with the pathogenesis of major depressive disorder (MDD) and generalized anxiety disorders (GAD)². This is due to the fact that selective serotonin reuptake inhibitors (SSRIs) are effective pharmacological treatment for these psychiatric disorders^3,4. In addition, accumulative evidences suggest that mania⁵ and suicidal behavior⁶ may be associated with lower levels of serotonin functioning in the DRN. It has also been reported that Pet1-Cre;Lmx1b^flox/flox mice and hTM-DTA^iPet1 mice (genetic mouse models lacking most central serotonergic neurons from late embryonic stage⁷ and adulthood⁸, respectively) display enhanced contextual fear memory. However, despite extensive research, the exact involvement of DRN serotonergic neurons in these psychiatric disorders remains to be elucidated.

In order to explore the mechanisms by which DRN serotonergic neurons regulate the pathogenesis of the serotonin-associated psychiatric disorders, animal models have been generated. Optogenetic tools have been applied to inhibit serotonergic neurons in rat DRN, and these animals display increased anxiety-like behaviors⁹. However, optogenetics has limitations. For example, a light-delivery device must be implanted into the targeted region deep within the brain, and the surrounding tissue may be injured during implantation surgery or by heat emitted from the light device. Even if temperature alteration may not cause detectable brain tissue damage, it can still induce remarkable physiological and behavioral effects¹⁰.

Pharmacological manipulation may be an easier approach to create DRN serotonergic neuron-lesioned animal models. Some groups have generated DRN serotonergic neuron-lesioned rats by stereotaxic microinjection of serotonin neurotoxin 5,7-DHT in the DRN. However, these rat models display different behavioral alterations, such as anxiolytic behavior¹¹, increased anxiety-like behavior¹², and impaired object memory¹³. Despite many studies in rats, fewer studies have been performed on the influences of 5,7-DHT on mice. One group reported excessive mortality (>50%) and limited serotonin depletion in experimental mice that received stereotaxic microinjections of 5,7-DHT in the DRN¹⁴. Another group reported that unpredictable chronic mild stress (UCMS) can induce significant attack latency alteration in 5,7-DHT-induced DRN-lesioned mice. However, no histological results were provided to confirm the exact serotonergic neuron loss in the DRN¹⁵. Stereotaxic injection in the DRN using standard procedures may lead to massive bleeding and high mortality to mice, given the fact that the anatomical location of DRN is below the SSS¹⁶.

This protocol describes the protocol to generate a DRN serotonergic neuron-lesioned mouse model (>90% survival rate of the experimental mice) with stable loss of DRN serotonergic neurons by stereotaxic injection of 5,7-DHT. The injection in DRN uses an angled approach to prevent the injury to the SSS. This surgery consistently causes >70% loss of serotonergic neuron in the DRN of mice, and it produces anxiety-associated behavior alterations. The protocol used here is for inducing DRN lesions, but it can also be useful to researchers who want to perform stereotaxic injections in other midline structures. In addition, this DRN serotonergic neuron-lesioned mouse model provides a valuable tool for understanding the role of serotonergic neurons in psychiatric disorders (i.e., MDD and GAD) and assessing potential neuroprotective agents or therapeutic strategies for these conditions.

Protocol

All surgical interventions and animal care procedures have been approved by the Animal Committee of School of Life Sciences and Technology, Tongji University, Shanghai, China.

1. Housing of animals

1. Maintain male C57BL/6NCrl mice (10 weeks old, 25 g, n = 21) in standard conditions (24 °C temperature; 55% humidity) under a 12 h/12 h light/dark cycle.
2. Provide food and water ad libitum.
   NOTE: Here, three of the mice are used for confirming the needle track.

2. Preparation of reagents

NOTE: All drug preparation steps must be performed in a laminar flow hood to avoid contamination. All prepared solutions are stored in a -80 °C freezer. It is recommended to use one aliquot at a time, thaw completely, mix well before use, discard the leftovers in the tube, and avoid repeated freezing and thawing.

1. Dissolve 0.25 g of desipramine hydrochloride in 100 mL of 0.9% saline to yield a 2.5 mg/mL solution. Sterilize the solution using a syringe filter with 0.22 μm pore size hydrophilic PES membrane. Then, aliquot the solution (1.8 mL/tube) in 2 mL microcentrifuge (EP) tubes. Label, date, and store in a -80 °C freezer immediately. The recommended dosage for animal use is 25 mg/kg.
2. Dissolve 5 mg of 5,7-DHT in 1.67 mL of 0.9% saline containing 0.1% ascorbic acid to yield a 3 μg/μL solution. Vortex gently to mix, sterilize the solution using a syringe filter (0.22 μm pore size), and aliquot the solution (10 μL/tube) in 0.5 mL EP tubes. Label, date, and freeze at -80 °C immediately. The recommended dosage for animal use is 2 μL/mouse.
3. Dissolve ketoprofen (analgesic) following the previously described method¹⁷. Dissolve 250 mg of ketoprofen in 15 mL of water and 1 mL of 1 M NaOH, adjust the pH to 7.3 and make up the final volume to 125 mL, which will result in a final concentration of 2 mg/mL. Sterilize the solution using a 0.22 μm syringe filter and aliquot the solution (0.4 mL/tube) in 1.5 mL EP tubes. Label, date, and freeze at -80 °C until use. The recommended dosage for animal use is 5 mg/kg.

3. Preparation of instruments and mice

1. Prepare a surgical pack (containing a scalpel, pair of tissue forceps, pair of scissors, needle holder, 3-0 sutures, ear tags, and ear tag applicator) previously sterilized in high temperature autoclaves. Place 75% ethanol, povidone-iodine, 3% hydrogen peroxide, vehicle solution (0.1% ascorbic acid in 0.9% saline), a cotton swab, ofloxacin eye ointment, and a mouse recovery cage on top of a heating pad, using a Hamilton syringe with a 32 G needle and 1 cc syringe.
2. h prior to 5,7-DHT injection, weigh and record body weights of the mice, then subject them to intraperitoneal (i.p.) injections of desipramine (2.5 mg/mL, 10 μL/g weight).
   NOTE: The purpose of desipramine administration 1 h prior to 5,7-DHT injection is to prevent catacholaminergic cell loss¹⁸.
3. Anesthetize mice using isoflurane inhalation anesthesia (3% during induction, 1.5% during maintenance, flow rate = 2 L/min). Administer ketoprofen (2 mg/mL, 2.5 μL/g weight) via subcutaneous (s.c.) injection. Confirm adequate anesthesia depth by the absence of tail pinch response.

4. Stereotaxic injection

1. Place the anesthetized mouse on the stereotaxic platform, then fix its head with the ear bars and incisor bar of the stereotaxic apparatus.
2. Shave the mouse's head, then clean the exposed scalp with one scrub of 75% ethanol followed by one scrub of povidone-iodine.
3. Apply lidocaine ointment on the scalp using a cotton swab to provide local analgesia. Put ofloxacin eye ointment on the eyes to protect the cornea.
4. Make an incision on the scalp along the midline using a scalpel from 1 mm posterior to the eyes to the interaural line.
5. Use a 3% hydrogen peroxide-soaked swab to remove the periosteum, then dry the skull and expose the cranial sutures. Mark the location of the bregma and lambda.
6. Adjust the head position using the incisor bar until the bregma and lambda lay in the same horizontal plane. Adjust the needle tip to touch the bregma or lambda, record the medial/lateral (ML) and dorsal/ventral (DV) coordinates, and adjust the incisor bar so that the DV and ML coordinates of bregma and lambda are equal, respectively.
7. Unlock the perpendicular positioning button and lock screw, then set the manipulator arm (z-axis) to 30° in the anterior/posterior (AP) direction as shown in Figure 1A,B. Lock the button.
8. Fix the Hamilton syringe filled with 2 μL of 5,7-DHT (3 μg/μL) onto the holder (for the sham group, mice are injected with 0.9% saline containing 0.1% ascorbic acid). Adjust the needle tip to touch the bregma landmark, then zero the ML, AP, and DV values using the digital display module.
   NOTE: The 5,7-DHT solution is brown colored liquid, making it easy to determine whether the Hamilton syringe is properly filled. If there is no access to a digital display module, record the number displayed on three axes when the needle tip touches the bregma, and the final coordinates represent “the recorded number plus the coordinate provided in this protocol”. For example, if the recorded number on the y-axis (A/P direction) manipulator arm is “a”, then the final coordinate in the A/P direction should be “a - 6.27”.
9. Move the manipulator arm to adjust the needle tip to the injection position (APa = -6.27, ML = 0; formula for calculating the final coordinates is listed in Figure 1B). Mark the target position using a marker pen.
10. Drill the burr hole using portable micromotor high-speed drill, then move the manipulator arm to lower the needle tip to the target (DVa = -4.04). Inject 2 μL of the solution into the brain slowly (0.5 μL/3 min), keeping the needle in situ for an additional 5 min to prevent solution leakage. Remove the needle gently after injection.
11. Apply interrupted sutures over the incision using 3-0 sutures. Wrap the sutured incision with a cotton swab soaked in povidone-iodine to avoid infection.
12. Clean the right ear with a povidone-iodine cotton swab, then apply the sterilized ear tag to the base of the ear for identification.
13. Remove experimental mice from the stereotaxic apparatus and place in a mice recovery cage on top of a heating pad until full recovery from anesthesia is observed.

5. Postoperative care of mice

1. Subject mice to s.c. injections of ketoprofen 1x/day up to 2 days after surgery.
2. Inspect mice daily up to 7 days after surgery.

6. Elevated T-maze test

NOTE: Perform the test as described previously^19,20.

1. Ensure that the elevated T-maze (ETM) test is comprised of two open arms (30 cm x 5 cm x 0.5 cm) perpendicular to two enclosed arms (30 cm x 5 cm x 16 cm x 0.5 cm) with a center platform (5.0 cm x 5.0 cm x 0.5 cm). One of the enclosed arms is blocked by an opaque plastic sheet to form a T-shape (Figure 2A).
2. Three days before the ETM test, habituate the animals by handling daily for 5 min.
3. On day 30 after surgery, perform the ETM test.
4. On the day of behavioral testing, expose mice to one of the open arms for 10 min.
5. For inhibitory avoidance testing, place each mouse in the distal end of the enclosed arm and record the time taken to leave this arm with four paws in three trials. The first trial is baseline avoidance, the second trial is avoidance 1, and the third trial is avoidance 2.
6. Keep mice in their cages for 30 s between trails and establish a cutoff time (here, 300 s is used for each trial).

7. Perfusion, fixation, immunohistochemical staining, and quantification

1. At the end of the study (35 days after surgery, after the behavioral test), perform whole body perfusion and fixation once the mouse is under general anesthesia.
   1. Anesthetize the mouse as described in step 3.3. Place the deeply anesthetized mouse in dorsal recumbency.
   2. Wet the fur with 75% alcohol over the entire ventral area.
   3. Make a cut below the sternum followed by a V-shape incision in the rib cage. Grasp the rib cage using the clamp to expose the heart.
   4. Insert the venous infusion needle into the left ventricle, then cut the atrial appendage immediately with scissors to let the blood flow out.
   5. Turn on the peristaltic pump to perfuse saline into the whole body through the circulatory system. Switch from saline to 4% paraformaldehyde (PFA) when fluid exiting the right atrium becomes clear and when the liver changes from red to pale red in color. Perfuse another 5 mL of 4% PFA when the mouse tail moves, and the body is stiff.
      NOTE: Swaying of the mouse tail is the sign of adequate perfusion.
2. Isolation of the brain after intracardiac perfusion with 4% PFA
   1. Decapitate the mouse using large scissors, then cut the skin to expose the skull.
   2. Make two cuts at the base of the skull (connected to the neck), make one cut along the line linking the eyes, then cut the skull along the sagittal suture. Grasp and peel the skull of each hemisphere outward to expose the brain using forceps.
      NOTE: Cutting the skull along the sagittal suture must be performed carefully, otherwise the brain may be damaged and separate into parts.
   3. Remove the brain and place it in 4% PFA at 4 °C overnight for post-fixation, then transfer the brain into 30% sucrose until it sinks to the bottom.
      NOTE: Dehydration using 30% sucrose is not necessary if sections are cut with a vibratome.
3. Absorb any excess liquid around the tissue using a paper towel prior to embedding. Embed the brain tissue in OCT with the rostral face on the chuck.
   NOTE: Orientation of the sample is important. The rostral face of the brain must be attached to the chuck to make sure that the cutting edge is the caudal portion.
4. Cut a coronal section of the DRN (4.0–4.8 mm posterior from bregma) at 30 μm thickness (four-section intervals) using a cryostat and collect the sections in PBS. Store in brain section cryopreservation solution (SCS) at -20 °C.
   NOTE: Identification of the DRN is very important. The consecutive anatomical structures are shown as Figure 3A–H. The characteristic structures are as follows: lateral recess of the fourth ventricle (LR4V, red dash line in Figure 3A,E), fourth ventricle (4V, red dash triangle in Figure 3B,F), second cerebellar lobule (2Cb, red dash diamond-shape structure in Figure 3C/G), and aqueduct (Aq, red dash hole in Figure 3D,H). Start to collect the tissues when structures are observed as shown in Figure 3D,H, then cut in 30 μm thick sections (four-section intervals) to yield 24 total coronal sections. H&E staining of the sections is shown in (Figure 3E–H). The fourth section of each of the four sections will be selected to perform immunofluorescent staining. The remaining sections are stored in the SCS. If it is difficult to recognize the structures, mounting one piece of the sections on the slide is helpful.
5. Perform immunohistochemical assay as described previously^21,22.
6. Count 5-HT-positive cells at different section levels throughout the entire DRN using ImageJ.

8. Statistical analysis

1. Use statistical software for data analysis. Express the results as means ± SEM.
2. Process the values for statistical analysis by two-way ANOVA or unpaired t-test and consider the differences significant at **p < 0.01.

Results

In a coronal section, the location of the DRN is just below the SSS and aqueduct (Figure 1B,C); thus, targeting the DRN using standard procedures can lead to massive bleeding and high mortality in mice¹⁶. Therefore, stereotaxic injections were performed here using an angled approach instead of the standard vertical approach to avoid damage to the SSS (Figure 1A,B). To confirm the location of the needle en...

Discussion

This protocol successfully describes production of a reliable DRN serotonergic neuron-lesioned mouse model with high lesion reproducibility and low mortality rate. Targeting the DRN is a complex task, since it can damage the SSS located just above the DRN¹⁶ and lead to excessive bleeding and even death¹⁴. Therefore, stereotaxic injections were performed by setting the manipulation arm at 30° in the AP direction to avoid injury to the SSS (Figure 1<...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.22micron syringe filter	Millipore	SLGPRB
3% hydrogen peroxide	Caoshanhu Co.,Ltd, Jiangxi, China
5,7-Dihydroxytryptamine	Sigma-Aldrich	SML2058	3ug/ul, 2ul
Compact small animal anesthesia machine	RWD Life Science Co., Ltd	R500 series
Cryostat	Leica Biosystems, Wetzlar, Germany	CM1950
Cy 3 AffiniPure Donkey Anti-Goat IgG (H+L)	Jackson ImmunoResearch	705-165-003	1:2,000
dapi	Sigma-Aldrich	D8417
desipramine hydrochloride	Sigma-Aldrich	PHR1723	25mg/kg
Eppendorf tube	Quality Scientific Plastics	509-GRD-Q
goat anti-5-HT antibody	Abcam	ab66047	1:800
GraphPad Prism	Graphpad Software Inc, CA, US
Hamilton Microliter syringe	Hamilton	87943
Ketoprofen	Sigma-Aldrich	K1751-1G	5mg/kg
L-ascorbic acid	BBI Life Sciences	A610021-0500	0.10%
lidocaine ointment	Tsinghua Tongfang Pharmaceutical Co. Ltd	H20063466
ofloxacin eye ointment	Shenyang Xingqi Pharmaceutical Co.Ltd, China	H10940177
peristaltic pump	Huxi Analytical Instrument Factory Co., Ltd, Shanghai, China	HL-1D
stereotaxic apparatus	RWD Life Science Co., Ltd	68018
ultra-low temperature freezer	Haier	DW-86L388
Vortex	Kylin-bell	VORTEX-5