This study was supported by the National Health and Medical Research Council of Australia (NHMRC) Project grant 1022839. AJH is an Australian Research Council (ARC) FT3 Future Fellow (FT100100835). The Florey Institute of Neuroscience and Mental Health acknowledges support from the Victorian Government&#8217;s Operational Infrastructure Support Grant.

NOTE: All experimental procedures on animals were approved by the Florey Institute of Neuroscience &#38; Mental Health Animal Ethics Committee and conform to Australia&#8217;s National Health and Medical Research Council published code of practice for the care and use of animals for scientific purposes (7th edition, 2004).
1. Environmental Manipulations
<ol>
	<li>Gender Pairing
	<ol>
		<li>Use sexually mature (&#62;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).</li>
		<li>Upon arrival into the animal holding facility, group-house the mice by gender for &#62;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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Environmental Enrichment
	<ol>
		<li>Use sexually mature (&#62;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.</li>
		<li>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 &#62;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.</li>
		<li>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.
		<ol>
			<li>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.</li>
			<li>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.</li>
		</ol>
		</li>
		<li>Subject EE mice to additional &#8216;super-enrichment&#8217; (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.
		<ol>
			<li>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.</li>
			<li>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).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

2. Osmotic Pump and Brain Infusion Cannula Implants for Drug Infusion
<ol>
	<li>Preparation
	<ol>
		<li>Use sterile osmotic pumps and brain infusion kits, which are available commercially. The day before implantation, using aseptic&#160;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 &#176;C. Incubate drug- and vehicle-filled implants separately to avoid cross-contamination.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>Sterilize all surgical equipment and employ aseptic&#160;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&#8217;s body temperature remains normal throughout the procedure.</li>
		<li>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 &#8216;pocket&#8217; 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.</li>
		<li>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.</li>
		<li>With the cannula held by a cannula holder, place the pump and connecting tube into the subcutaneous &#8216;pocket&#8217; overlying the mouse&#8217;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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Post-surgical Treatment of Animals
	<ol>
		<li>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.</li>
		<li>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 &#62;10% body weight, social withdrawal, lack of grooming, &#8220;fluffing&#8221;, movement dysfunction, seizures) with systemic analgesics and/or antibiotics as necessary.</li>
		<li>Euthanize mice if &#62;15% weight loss or symptoms of sickness, pain, stress or discomfort are non-recoverable within 1 week of remedial treatment.</li>
	</ol>
	</li>
</ol>
3. Brain Tissue Preparation, Immunohistochemical Processing, and Stereology
<ol>
	<li>Perfusion
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Clamp the cannula in place then pump warm (37 &#176;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 &#176;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.</li>
	</ol>
	</li>
	<li>Preparation of Free-floating Cryosections
	<ol>
		<li>Cut serial sections through the brain regions of interest and collect these in PBS. 
		NOTE: We cut 40 &#181;m thick sections using a cryostat. Other types of tissue sectioning might be applied depending on the expertise in the corresponding laboratory.</li>
	</ol>
	</li>
	<li>Immunohistochemistry
	<ol>
		<li>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 &#176;C for 48 hr, then polyclonal biotinylated goat anti-rabbit (1:1,000) at room temperature for 2 hr.</li>
		<li>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.</li>
		<li>Mount the sections on gelatinized microscope slides, air dry them, then Nissl stain (neutral red), dehydrate in alcohol, clear (X-3B), and coverslip.</li>
	</ol>
	</li>
	<li>Stereology
	<ol>
		<li>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 &#60;5 &#181;m and count only those cells with a visible nucleus.</li>
		<li>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 &#181;m = 3025 &#181;m2) at regular pre-determined intervals (x = 140 &#181;m, y = 140 &#181;m for SNc; x = 100 &#181;m, y = 100 &#181;m for VTA and LC) throughout each nucleus in every fourth section.</li>
	</ol>
	</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+environmental+enrichment+exposure+on+synaptic+transmission+and+plasticity+in+the+hippocampus.">Effects of environmental enrichment exposure on synaptic transmission and plasticity in the hippocampus.</a> Current topics in behavioral neurosciences. 15, 165-187 (2013).</li><li>Sztainberg, Y., Chen, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+environmental+enrichment+model+for+mice.">An environmental enrichment model for mice.</a> Nature protocols. 5, 1535-1539 (2010).</li><li>Sale, A., Berardi, N., Maffei, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Environment+and+brain+plasticity:+towards+an+endogenous+pharmacotherapy.">Environment and brain plasticity: towards an endogenous pharmacotherapy.</a> Physiological reviews. 94, 189-234 (2014).</li></ol>

The authors have nothing to disclose.

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...

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&#8217;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 administration2,3, 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 &#8216;DA phenotype&#8217; 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 cells4,5. 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 neurons4,5. 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 neurons4,5. 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.

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			<td>Merck Millipore Headquarters&#160;, 290 Concord road, Billerica, MA 01821</td>
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			<td>MicroBrightField Bioscience, 185 Allen Brook Lane, Suite 101, Williston, VT 05495</td>
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environmental modulations of the number of midbrain dopamine neurons in adult mice

Long-lasting changes in the brain or &#8216;brain plasticity&#8217; 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 &#8216;enriched environments&#8217; (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 &#8216;DA&#8217; 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&#8217;s disease, attention deficit and hyperactivity disorder, schizophrenia, and drug addiction).

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 (GABAA 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...

Watch this Scientific Journal Video about Environmental Modulations of the Number of Midbrain Dopamine Neurons in Adult Mice at JoVE.com

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

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.

Long-lasting changes in the brain or &#8216;brain plasticity&#8217; underlie adaptive behavior and brain repair following disease or injury. Furthermore, ...

environmental-modulations-number-midbrain-dopamine-neurons-adult

Research

JoVE Journal

Neuroscience

8.7K Views.  The University of Melbourne. 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.

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

Application of a NMDA Receptor Conductance in Rat Midbrain Dopaminergic Neurons Using the Dynamic Clamp Technique

Neuroscientists study the function of the brain by investigating how neurons in the brain communicate. Many investigators look at changes in the electrical activity of one or more neurons in response to an experimentally-controlled input. The electrical activity of neurons can be recorded in isolated brain slices using patch clamp techniques with glass micropipettes. Traditionally, experimenters can mimic neuronal input by direct injection of current through the pipette, electrical stimulation of the other cells or remaining axonal connections in the slice, or pharmacological manipulation by receptors located on the neuronal membrane of the recorded cell.
Direct current injection has the advantages of passing a predetermined current waveform with high temporal precision at the site of the recording (usually the soma). However, it does not change the resistance of the neuronal membrane as no ion channels are physically opened. Current injection usually employs rectangular pulses and thus does not model the kinetics of ion channels. Finally, current injection cannot mimic the chemical changes in the cell that occurs with the opening of ion channels.
Receptors can be physically activated by electrical or pharmacological stimulation. The experimenter has good temporal precision of receptor activation with electrical stimulation of the slice. However, there is limited spatial precision of receptor activation and the exact nature of what is activated upon stimulation is unknown. This latter problem can be partially alleviated by specific pharmacological agents. Unfortunately, the time course of activation of pharmacological agents is typically slow and the spatial precision of inputs onto the recorded cell is unknown.
The dynamic clamp technique allows an experimenter to change the current passed directly into the cell based on real-time feedback of the membrane potential of the cell (Robinson and Kawai 1993, Sharp et al., 1993a,b; for review, see Prinz et al. 2004). This allows an experimenter to mimic the electrical changes that occur at the site of the recording in response to activation of a receptor. Real-time changes in applied current are determined by a mathematical equation implemented in hardware.
We have recently used the dynamic clamp technique to investigate the generation of bursts of action potentials by phasic activation of NMDA receptors in dopaminergic neurons of the substantia nigra pars compacta (Deister et al., 2009; Lobb et al., 2010). In this video, we demonstrate the procedures needed to apply a NMDA receptor conductance into a dopaminergic neuron.

In this video, we demonstrate how to apply a conductance into a dopaminergic neuron recorded in the whole cell configuration in rat brain slices. This technique is called the dynamic clamp.

Neuroscientists study the function of the brain by investigating how neurons in the brain communicate. Many investigators look at changes in the ...

Studying the Integration of Adult-born Neurons

Neurogenesis occurs in adult mammalian brains in the sub-ventricular zone (SVZ) of the lateral ventricle and in the sub-granular zone (SGZ) of the hippocampal dentate gyrus throughout life. Previous reports have shown that adult hippocampal neurogenesis is associated with diverse brain disorders, including epilepsy, schizophrenia, depression and anxiety (1). Deciphering the process of normal and aberrant adult-born neuron integration may shed light on the etiology of these diseases and inform the development of new therapies.
SGZ adult neurogenesis mirrors embryonic and post-natal neuronal development, including stages of fate specification, migration, synaptic integration, and maturation. However, full integration occurs over a prolonged, 6-week period. Initial synaptic input to adult-born SGZ dentate granule cells (DGCs) is GABAergic, followed by glutamatergic input at 14 days (2). The specific factors which regulate circuit formation of adult-born neurons in the dentate gyrus are currently unknown.
Our laboratory uses a replication-deficient retroviral vector based on the Moloney murine leukemia virus to deliver fluorescent proteins and hypothesized regulatory genes to these proliferating cells. This viral technique provides high specificity and resolution for analysis of cell birth date, lineage, morphology, and synaptogenesis.
A typical experiment often employs two or three viruses containing unique label, transgene, and promoter elements for single-cell analysis of a desired developmental process in vivo. The following protocol describes a method for analyzing functional newborn neuron integration using a single green (GFP) or red (dTomato) fluorescent protein retrovirus and patch-clamp electrophysiology.

A way to study the integration of newborn dentate granule cells in adult animals is described. This technique uses an engineered retrovirus to label newborn neurons, followed by electrophysiological recordings to determine in vivo functional integration.

Neurogenesis occurs in adult mammalian brains in the sub-ventricular zone (SVZ) of the lateral ventricle and in the sub-granular zone (SGZ) of the ...

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls gastrointestinal motility. A procedure for the isolation and culture of a mixed population of neurons and glia from the myenteric plexus is described. The primary cultures can be maintained for over 7 days, with connections developing among the neurons and glia. The longitudinal muscle strip with the attached myenteric plexus is stripped from the underlying circular muscle of the mouse ileum or colon and subjected to enzymatic digestion. In sterile conditions, the isolated neuronal and glia population are preserved within the pellet following centrifugation and plated on coverslips. Within 24-48 hr, neurite outgrowth occurs and neurons can be identified by pan-neuronal markers. After two days in culture, isolated neurons fire action potentials as observed by patch clamp studies. Furthermore, enteric glia can also be identified by GFAP staining. A network of neurons and glia in close apposition forms within 5 - 7 days. Enteric neurons can be individually and directly studied using methods such as immunohistochemistry, electrophysiology, calcium imaging, and single-cell PCR. Furthermore, this procedure can be performed in genetically modified animals. This methodology is simple to perform and inexpensive. Overall, this protocol exposes the components of the enteric nervous system in an easily manipulated manner so that we may better discover the functionality of the ENS in normal and disease states.

An In-vitro Preparation of Isolated Enteric Neurons and Glia from the Myenteric Plexus of the Adult Mouse

We demonstrate a cell culture protocol for the direct study of neuronal and glial components of the enteric nervous system. A neuron/glia mixed culture on coverslips is prepared from the myenteric plexus of adult mouse providing the ability to examine individual neuron and glia function by electrophysiology, immunohistochemical, etc.

The enteric nervous system is a vast network of neurons and glia running the length of the gastrointestinal tract that functionally controls ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

Laser Capture Microdissection - A Demonstration of the Isolation of Individual Dopamine Neurons and the Entire Ventral Tegmental Area

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue sections on a microscope slide. When combined with immunohistochemistry, LCM can be used to isolate individual cells types based on a specific protein marker. Here, the LCM technique is described for collecting a specific population of dopamine neurons directly labeled with&#160;tyrosine hydroxylase immunohistochemistry and for isolation of the dopamine neuron containing region of the ventral tegmental area using indirect tyrosine hydroxylase immunohistochemistry on a section adjacent to those used for LCM. An infrared (IR) capture laser is used to both dissect individual neurons as well as the ventral tegmental area off glass slides and onto an LCM cap for analysis. Complete dehydration of the tissue with 100% ethanol and xylene is critical. The combination of the IR capture laser and the ultraviolet (UV) cutting laser is used to isolate individual dopamine neurons or the ventral tegmental area when using PEN membrane slides. A PEN membrane slide has significant advantages over a glass slide as it offers better consistency in capturing and collecting cells, is faster collecting large pieces of tissue, is less reliant on dehydration and results in complete removal of the tissue from the slide. Although removal of large areas of tissue from a glass slide is feasible, it is considerably more time consuming and frequently leaves some residual tissue behind. Data shown here demonstrate that RNA of sufficient quantity and quality can be obtained using these procedures for quantitative PCR measurements. Although RNA and DNA are the most commonly isolated molecules from tissue and cells collected with LCM, isolation and measurement of microRNA, protein and epigenetic changes in DNA can also benefit from the enhanced anatomical and cellular resolution obtained using LCM.

The isolation of individual dopamine neurons or the ventral tegmental area with direct or indirect immunohistochemistry is demonstrated using laser capture microdissection. Parameters for isolation of tissue from a glass slide using an infrared laser and from membrane slides using the combination of an infrared and ultraviolet laser are discussed.

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...

Quantification of Endosome and Lysosome Motilities in Cultured Neurons Using Fluorescent Probes

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, survival, and glial communications. To date, numerous studies have reported that defects in these systems cause various neuronal diseases. Thus, understanding the mechanisms underlying vesicle dynamics may provide influential clues that could aid in the treatment of several neuronal disorders. Here, we describe a method for quantifying vesicle motilities, such as motility distance and rate of movement, using a software plug-in for the ImageJ platform. To obtain images for quantification, we labeled neuronal endosome-lysosome structures with EGFP-tagged vesicle marker proteins and observed the movement of vesicles using a time-lapse microscopy. This method is highly useful and simplify measuring vesicle motility in neurites, such as axons and dendrites, as well as in the soma of both neurons and glial cells. Furthermore, this method can be applied to other cell lines, such as fibroblasts and endothelial cells. This approach could provide a valuable advancement of our understanding of membrane trafficking.

The investigation of membrane trafficking is crucial for understanding neuronal functions. Here, we introduce a method for quantifying vesicle motility in neurons. This is a convenient method that can be adapted to the quantification of membrane trafficking in the nervous system.

In the brain, membrane trafficking systems play important roles in regulating neuronal functions, such as neuronal morphology, synaptic plasticity, ...

Assessment of Dopaminergic Homeostasis in Mice by Use of High-performance Liquid Chromatography Analysis and Synaptosomal Dopamine Uptake

Dopamine (DA) is a modulatory neurotransmitter controlling motor activity, reward processes and cognitive function. Impairment of dopaminergic (DAergic) neurotransmission is strongly associated with several central nervous system-associated diseases such as Parkinson&#39;s disease, attention-deficit-hyperactivity disorder and drug addiction1,2,3,4. Delineating disease mechanisms involving DA imbalance is critically dependent on animal models to mimic aspects of the diseases, and thus protocols that assess specific parts of the DA homeostasis are important to provide novel insights and possible therapeutic targets for these diseases.
Here, we present two useful experimental protocols that when combined provide a functional read-out of the DAergic system in mice. Biochemical and functional parameters on DA homeostasis are obtained through assessment of DA levels and dopamine transporter (DAT) functionality5. When investigating the DA system, the ability to reliably measure endogenous levels of DA from adult brain is essential. Therefore, we present how to perform high-performance liquid chromatography (HPLC) on brain tissue from mice to determine levels of DA. We perform the experiment on tissue from dorsal striatum (dStr) and nucleus accumbens (NAc), but the method is also suitable for other DA-innervated brain areas.
DAT is essential for reuptake of DA into the presynaptic terminal, thereby controlling the temporal and spatial activity of released DA. Knowing the levels and functionality of DAT in the striatum is of major importance when assessing DA homeostasis. Here, we provide a protocol that allows to simultaneously deduce information on surface levels and function using a synaptosomal6 DA uptake assay.
Current methods combined with standard immunoblotting protocols provide the researcher with relevant tools to characterize the DAergic system.

Synaptosomal dopamine uptake and high-performance liquid chromatography analysis represent experimental tools to investigate dopamine homeostasis in mice by assessing the function of the dopamine transporter and levels of dopamine in striatal tissue, respectively. Here we present protocols to measure dopamine tissue content and assess the functionality of the dopamine transporter.

Dopamine (DA) is a modulatory neurotransmitter controlling motor activity, reward processes and cognitive function. Impairment of dopaminergic (DAergic) ...

Use of Synaptic Zinc Histochemistry to Reveal Different Regions and Laminae in the Developing and Adult Brain

Characterization of anatomical and functional brain organization and development requires accurate identification of distinct neural circuits and regions in the immature and adult brain. Here we describe a zinc histochemical staining procedure that reveals differences in staining patterns among different layers and brain regions. Others have utilized this procedure not only to reveal the distribution of zinc-containing neurons and circuits in the brain, but also to successfully delineate areal and laminar boundaries in the developing and adult brain in several species. Here we illustrate this staining procedure with images from developing and adult ferret brains. We reveal a zinc-staining pattern that serves as an anatomical marker of areas and layers, and can be reliably used to distinguish visual cortical areas in the developing and adult visual cortex. The main goal of this protocol is to present a histochemical method that allows the accurate identification of layers and regions in the developing and adult brain where other methods fail to do so. Secondarily, in conjunction with densitometric image analysis, this method allows one to assess the distribution of synaptic zinc to reveal potential changes throughout development. This protocol describes in detail the reagents, tools, and steps necessary to successively stain frozen brain sections. Although this protocol is described using ferret brain tissue, it can easily be adapted for use in rodents, cats, or monkeys as well as in other brain regions.

We describe a histochemical procedure that reveals characteristic laminar and areal zinc staining patterns in different brain regions. The zinc-staining pattern may be used in conjunction with other anatomical markers to reliably distinguish layers and regions in the developing and adult brain.

Characterization of anatomical and functional brain organization and development requires accurate identification of distinct neural circuits and regions ...

Retrograde Neuroanatomical Tracing of Phrenic Motor Neurons in Mice

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic nerves innervating the respiratory diaphragm. In spinal cord slices, phrenic motor neurons cannot be identified from other motor neurons on morphological or biochemical criteria. We provide the description of procedures for visualizing phrenic motor neuron cell bodies in mice, following intrapleural injections of cholera toxin subunit beta (CTB) conjugated to a fluorophore. This fluorescent neuroanatomical tracer has the ability to be caught up at the diaphragm neuromuscular junction, be carried retrogradely along the phrenic axons and reach the phrenic cell bodies. Two methodological approaches of intrapleural CTB delivery are compared: transdiaphragmatic versus transthoracic injections. Both approaches are successful and result in similar number of CTB-labeled phrenic motor neurons. In conclusion, these techniques can be applied to visualize or quantify the phrenic motor neurons in various experimental studies such as those focused on the diaphragm-phrenic circuitry.

Here, we describe a protocol for identifying phrenic motor neurons in mice after intrapleural delivery of fluorophore conjugated cholera toxin subunit beta. Two techniques are compared to inject the pleural cavity: transdiaphragmatic versus transthoracic approaches.

Phrenic motor neurons are cervical motor neurons originating from C3 to C6 levels in most mammalian species. Axonal projections converge into phrenic ...