Environmental Modulations of the Number of Midbrain Dopamine Neurons in Adult Mice

Summary

This protocol describes two different environmental manipulations and a concurrent brain infusion protocol to study environmentally-induced brain changes underlying adaptive behavior and brain repair in adult mice.

Abstract

Long-lasting changes in the brain or ‘brain plasticity’ underlie adaptive behavior and brain repair following disease or injury. Furthermore, interactions with our environment can induce brain plasticity. Increasingly, research is trying to identify which environments stimulate brain plasticity beneficial for treating brain and behavioral disorders. Two environmental manipulations are described which increase or decrease the number of tyrosine hydroxylase immunopositive (TH+, the rate-limiting enzyme in dopamine (DA) synthesis) neurons in the adult mouse midbrain. The first comprises pairing male and female mice together continuously for 1 week, which increases midbrain TH+ neurons by approximately 12% in males, but decreases midbrain TH+ neurons by approximately 12% in females. The second comprises housing mice continuously for 2 weeks in ‘enriched environments’ (EE) containing running wheels, toys, ropes, nesting material, etc., which increases midbrain TH+ neurons by approximately 14% in males. Additionally, a protocol is described for concurrently infusing drugs directly into the midbrain during these environmental manipulations to help identify mechanisms underlying environmentally-induced brain plasticity. For example, EE-induction of more midbrain TH+ neurons is abolished by concurrent blockade of synaptic input onto midbrain neurons. Together, these data indicate that information about the environment is relayed via synaptic input to midbrain neurons to switch on or off expression of ‘DA’ genes. Thus, appropriate environmental stimulation, or drug targeting of the underlying mechanisms, might be helpful for treating brain and behavioral disorders associated with imbalances in midbrain DA (e.g. Parkinson’s disease, attention deficit and hyperactivity disorder, schizophrenia, and drug addiction).

Introduction

DArgic signaling by neurons in the ventral tegmental area (VTA) and substantia nigra pars compacta (SNc) of the midbrain is thought to be important for reward-motivated cognitive, emotive and motor behaviors. However, too much or too little midbrain DA signaling causes many disabling symptoms in a variety of neurological disorders (e.g. Parkinson’s disease, attention deficit and hyperactivity disorder, schizophrenia, and drug addiction). Drugs that increase or decrease DA signaling alleviate these symptoms, however they also produce side-effects attributable to dysregulated signaling and off-target effects. Drug efficacy also declines over time due to compensatory responses of the brain. The challenge therefore is to restore normal midbrain DA signaling in a more targeted and physiological way, and a favored approach is by increasing or decreasing the number of midbrain DA neurons.

Evidence has been accumulating for several decades that the expression of genes and proteins involved in metabolizing and trafficking DA and other catecholamines in mature adult cells is modifiable (reviewed in¹). In midbrain, the number of tyrosine hydroxylase immunopositive (TH+, the rate-limiting enzyme in DA synthesis) neurons decreases then increases following neurotoxin administration^2,3, while the number of TH immunonegative (TH-) neurons shows the opposite pattern (i.e. increases then decreases³). This is consistent with loss then gain of the ‘DA phenotype’ by some cells. The number of TH+ and TH- SNc neurons has also been shown to change in equal but opposite directions following various treatments that alter the electrical activity of these cells^4,5. For example, infusion of the small-conductance, calcium-activated potassium (SK) channel antagonist apamin into midbrain for 2 weeks decreases the number of TH+ and increases (by the same amount) the number of TH- SNc neurons^4,5. In contrast, infusion of the SK channel agonist 1-EBIO increases the number of TH+ and decreases (by the same amount) the number of TH- SNc neurons^4,5. Similar changes were seen following a variety of treatments targeting SNc neuronal activity, including some which targeted afferent inputs⁴. This apparent regulation of the number of SNc DArgic neurons by neuronal activity and afferent input raises the possibility that the environment or behavior can influence the number of SNc neurons. Indeed adult mice exposed to different environments have more or less midbrain (SNc and VTA) TH+ neurons, and at least some of these environment-induced changes are abolished by concurrent blockade of synaptic input in midbrain⁶. The aims of this communication are to: (1) provide further details about how to implement our environmental manipulations and drug infusions; and (2) provide further data supporting our contention that the environment regulates the number of midbrain DA neurons, via afferent input.

Protocol

NOTE: All experimental procedures on animals were approved by the Florey Institute of Neuroscience & Mental Health Animal Ethics Committee and conform to Australia’s National Health and Medical Research Council published code of practice for the care and use of animals for scientific purposes (7^th edition, 2004).

1. Environmental Manipulations

1. Gender Pairing
   1. Use sexually mature (>8 week old), age-matched male and female mice.
      NOTE: We typically use C57BL/6 mice, but have had the same outcomes using Swiss mice in this protocol. We typically use n = 8 males and n = 8 females for each experiment, divided into n = 2 male-male pairs (n = 4 males), n = 2 female-female pairs (n = 4 females), and n = 4 male-female pairs (n = 4 males and n = 4 females).
   2. Upon arrival into the animal holding facility, group-house the mice by gender for >3 days to acclimatize. Here and throughout the gender pairing, keep extraneous environmental stimuli (e.g. ambient temperature, light:dark cycle, food, water, handling and cleaning) constant or distribute equally to all mice.
   3. Randomly assign each mouse to a male-male pair, a female-female pair, or a male-female pair. Place each pair into a clean cage in isolation (2 mice/cage) with ad libitum access to food and water, and simply house them in this way continuously for 7 days.
      NOTE: Routine animal husbandry is OK during this period but must be distributed equally to all mice. Take care to avoid pairing of male and female littermates.
2. Environmental Enrichment
   1. Use sexually mature (>8 week old), age-matched male or female mice.
      NOTE: We typically use n = 18 mice for each experiment, divided into n = 6/treatment group.
   2. Upon arrival into the animal holding facility keep them group-housed in identical standard housed (SH) (e.g. see below) conditions at this stage for >3 days to acclimatize. Here and throughout the experiment, keep extraneous environmental stimuli (e.g. ambient temperature, light:dark cycle, food, water, handling and cleaning) constant or distribute equally to all mice.
   3. To begin environmental enrichment, randomly assign each mouse to one of 3 groups: SH; running wheel (RW), or environment enriched (EE) and place each group of mice together into an identical clean cage with ad libitum access to food and water.
      NOTE: Here, larger rat-holding cages for all groups, measuring 27 cm wide, 42 cm long, and 16 cm deep were used.
      1. In addition, provide the following environmental conditions to each group: SH comprising only litter (paper pellets or sawdust) on the floor; RW comprising SH plus 2 running wheels; EE comprising RW plus toys (ropes, ladders, tunnels, and objects such as empty paper towel rolls and pieces of tissue paper) with which to explore, play, climb, hide, and nest.
      2. House them in this way continuously for 14 days.
         NOTE: Routine animal husbandry is OK during this period but must be distributed equally to all mice.
   4. Subject EE mice to additional ‘super-enrichment’ (SE) by placing them together into a larger cage containing novel toys for 1hour/day (same hour each day), 5days/week.
      NOTE: Here, a 46 cm wide, 69 cm long, and 40 cm deep plastic tub was used.
      1. Maintain novelty by presenting a different set of toys each session. Clean toys that are to be re-presented to mice with soapy water and 80% ethanol to remove scents.
      2. Following each SE session, return the mice to their EE cage. Handle SH and RW mice the same as SE mice (except for SE itself) throughout this period (e.g. remove and return each SH and RW mouse from and to its cage).

2. Osmotic Pump and Brain Infusion Cannula Implants for Drug Infusion

1. Preparation
   1. Use sterile osmotic pumps and brain infusion kits, which are available commercially. The day before implantation, using aseptic technique, prime the implants by filling each pump, connecting tube and cannula with sterile drug or vehicle solution (as described in the instructions supplied with the pumps). Connect them together and incubate in sterile saline overnight at 37 °C. Incubate drug- and vehicle-filled implants separately to avoid cross-contamination.
2. Surgery.
   NOTE: Depending on the country of the experimenter, special animal experimentation permits or allowances are required to pursue such types of surgery and post-operative experiments.
   1. Sterilize all surgical equipment and employ aseptic technique throughout. Anesthetize a mouse (e.g. using 1-2% isofluorane in air) and place it in a stereotaxic headframe. Check the depth of anesthesia throughout the procedure and administer further anesthetic as necessary (e.g. absence of limb withdrawal to noxious paw pinch is indicative of adequate anesthesia). Ensure the eyes are protected from desiccation by lubricant eye ointment, and that the mouse’s body temperature remains normal throughout the procedure.
   2. Make a midline incision through the skin starting from between the eyes and finishing 2 cm posterior to the back edge of the skull. Blunt dissect the skin away from the underlying fascia down the back of the mouse to create a subcutaneous ‘pocket’ large enough to comfortably fit the pump and connecting tube once the cannula is implanted (the connecting tube should not be bent upon completion). Scrape clear the fascia from over the dorsal aspect of the skull.
   3. Using an approximately 1.5 mm diameter dental burr, drill down into the skull at the appropriate stereotaxic coordinates until a thin, flexible layer of bone remains. Peel this layer away using fine forceps (this avoids damaging the underlying dura mater and brain with the dental burr). Ensure the skull is cleaned of any blood and bone fragments, and that there is no further bleeding.
   4. With the cannula held by a cannula holder, place the pump and connecting tube into the subcutaneous ‘pocket’ overlying the mouse’s back. Next, position the tip of the cannula at the appropriate stereotaxic coordinates and on the surface of the brain. Carefully lower the cannula into the brain but stop approximately 1 mm short of the required depth.
      NOTE: The remaining depth will be negotiated following application of the first layer of dental acrylic.
   5. Ensure again the surface of the skull is clean and dry, then prepare a small volume of dental acrylic and use a toothpick to smooth it over the entire exposed area of skull, including into the burr-hole through the skull around the cannula. Before the acrylic hardens, lower the cannula tip the remaining depth into the target.
   6. Prepare another small volume of dental acrylic and layer this over the first, as well as over the white plastic support for the cannula, to fix the cannula in place. Repeat with additional layers of acrylic as necessary.
   7. Once the acrylic has hardened, carefully remove the cannula holder and cut off the white plastic removable cannula tab using a scalpel blade heated with a butane flame. Lastly, suture the skin closed over the entire implant.
3. Post-surgical Treatment of Animals
   1. Apply antiseptic ointment to the skin margins, and administer an anti-inflammatory to the mouse (e.g. Meloxicam, 3 mg/kg s.c.). Remove the mouse from the frame, place it under a heat lamp, and observe until it regains consciousness. Do not return a mouse that has undergone surgery to the company of other animals until fully recovered.
      NOTE: Experimental manipulations such as gender pairing and environment enrichment can be initiated or resumed the day following surgery at the earliest.
   2. Monitor their body weight, general behavior and appearance daily. Treat signs of infection (e.g. swelling, redness, pus) around surgical incisions with topical antiseptic ointment. Treat any signs sickness, pain, stress or discomfort (e.g. loss of >10% body weight, social withdrawal, lack of grooming, “fluffing”, movement dysfunction, seizures) with systemic analgesics and/or antibiotics as necessary.
   3. Euthanize mice if >15% weight loss or symptoms of sickness, pain, stress or discomfort are non-recoverable within 1 week of remedial treatment.

3. Brain Tissue Preparation, Immunohistochemical Processing, and Stereology

1. Perfusion
   1. Immediately following environmental/drug manipulations, prepare the brain for study.
      NOTE: In the experiments reported here the number of SNc TH+ neurons were measured and compared in each of the different treatment groups.
   2. First administer an overdose of anesthetic to kill the mice (e.g. 100 mg/kg i.p. sodium pentobarbitone). Once anesthetized but before the heart stops beating, lay the mouse on its back and tie or pin down both forelimbs.
   3. Using a scalpel cut away the skin overlying the thorax then incise through the muscle just below the rib cage into the abdominal cavity. Place a clamp on the xiphoid process of the ribcage and lift the ribcage up and away from the liver exposing the diaphragm.
   4. Cut through the diaphragm and through the ribs laterally on both sides using large scissors until the ribcage can be folded back over the head to expose the lungs and heart. Using fine scissors make a small incision in the right atrium as an exit point for blood and perfused solutions, then cut through the base of the left ventricle and place a cannula up through the left ventricle, the left atrium, and into the aorta.
   5. Clamp the cannula in place then pump warm (37 °C) heparinized and 0.1 M phosphate buffered physiological saline (PBS) through the vasculature until the solution exiting the right atrium is entirely free of blood. Next, pump cold (4 °C) fixative solution (such as 4% paraformaldehyde in PBS) through the vasculature until the entire mouse is well fixed. Remove the brain and place in PBS with 30% sucrose for 2-3 days until the brain sinks.
      NOTE: Other types of tissue preparation might be applied depending on the expertise in the corresponding laboratory.
2. Preparation of Free-floating Cryosections
   1. Cut serial sections through the brain regions of interest and collect these in PBS.
      NOTE: We cut 40 µm thick sections using a cryostat. Other types of tissue sectioning might be applied depending on the expertise in the corresponding laboratory.
3. Immunohistochemistry
   1. Perform standard immunohistochemistry for proteins of interest. Incubate sections in 5% normal goat serum and 0.3% triton X-100 in PBS at room temperature for 30 min, then immunoreact with polyclonal rabbit anti-TH (1:400) at 4 °C for 48 hr, then polyclonal biotinylated goat anti-rabbit (1:1,000) at room temperature for 2 hr.
   2. Next, incubate in avidin-peroxidase (1:500) at room temperature for 1hr, then in cobalt- and nickel-intensified diamino-benzidine (0.5 mg/ml) at room temperature for 18-20 min; for the last 3-5 min of the diamino-benzidine incubation add hydrogen peroxide (0.01%) to catalyze chromagen precipitation. Wash sections three times for 10min each in PBS before, after, and between each of the above steps.
   3. Mount the sections on gelatinized microscope slides, air dry them, then Nissl stain (neutral red), dehydrate in alcohol, clear (X-3B), and coverslip.
4. Stereology
   1. Estimate the total number of TH+ and TH- SNc (and VTA and LC) neurons using unbiased stereological methods. Ensure that the stereologist is blind to the treatment received. Exclude glia on the basis of soma diameter <5 µm and count only those cells with a visible nucleus.
   2. Identify the SNc (and VTA and LC) by the spatial locations of TH+ cells and anatomical landmarks/boundaries according to the brain atlas of Paxinos and Watson⁷. Count TH+ cells within a counting frame (55 x 55 µm = 3025 µm²) at regular pre-determined intervals (x = 140 µm, y = 140 µm for SNc; x = 100 µm, y = 100 µm for VTA and LC) throughout each nucleus in every fourth section.

Results

Adult mice subjected to these environmental manipulations have altered numbers of midbrain (SNc and VTA), but not LC, TH+ neurons, and EE plus concurrent midbrain infusion of either picrotoxin or bicuculline (GABA_A receptor antagonists) abolishes EE-induction of more SNc TH+ neurons. These data were previously published in⁶. The present data were compiled in replicate experiments performed as part of that previous study, but have not been published elsewhere.

Specifically...

Discussion

Environmental manipulations

The motivation behind the design of these environmental manipulations (gender pairing and environmental enrichment) was to determine whether the environment, and/or behavior prompted by the environment, is associated with changes in the number of midbrain DA neurons. The focus was therefore on providing environments and stimulating behaviors that are likely to engage midbrain DA signaling. These included pairing with the opposite gender, and environmental e...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Isofluorane	Baxter Healthcare Pty Ltd, Baxter Drive, NSW 2146, Australia	AHN3640
ALZET Osmotic pump 1002	DURECT Corporation, PO Box 530 Cupertino, CA 95015-0530	0004317
ALZET Brain infusion kit 1	DURECT Corporation, PO Box 530 Cupertino, CA 95015-0530	0004760
ALZET cannula holder 1	DURECT Corporation, PO Box 530 Cupertino, CA 95015-0530	0008860
Vertex Monomer Self-curing (dental acrylic solvent)	Vertex Dental, Postbus 10, 3700 AA ZEIST, The Netherlands	n/a
Vertex Self Curing (dental acrylic powder)	Vertex Dental, Postbus 10, 3700 AA ZEIST, The Netherlands	n/a
METACAM (Meloxicam)	Troy Laboratories, 98 long Street, smithfield NSW 2164 Australia	L10100
Sodium Pentobarbitone	Lethabarb, Virbac, Milperra, NSW, Australia	571177
Normal goat serum	chemicon-temecula, CA	S26-Litre
Triton X-100	Merck Millipore Headquarters , 290 Concord road, Billerica, MA 01821	1.08603.1000
Polyclonal rabbit anti-tyrosine hydroxylase	Merck Millipore Headquarters , 290 Concord road, Billerica, MA 01821	AB152
Polyclonal biotinylated goat anti-rabbit	Dako Australia Pty. Ltd., Suite 4, Level 4, 56 Berry street, North Sydney, NSW, Australia 2060	EO432
Avidin peroxidase	Sigma-aldrich, Castle Hill, NSW 1765 AU	A3151-1mg
Diamino-benzidine	Sigma-aldrich, Castle Hill, NSW 1765 AU	D-5637
Stereo Investigator	MicroBrightField Bioscience, 185 Allen Brook Lane, Suite 101, Williston, VT 05495	n/a