This work was supported by the NIH/NIDDK (DK077106 to ECM).

1. Surgery Preparation
<ol>
 <li>Handling. Animals (most commonly, rats or mice, although the method for microdialysis and combination with behavioral testing is largely a general one with any required species-specific adaptations needed, e.g. for anesthesia) are handled for a minimum of 10 min/day for at least two days prior to surgery. Extensive handling has been shown to leave animals in an unstressed state at the time of testing, avoiding possible confound4-5,7,8. We will use rats throughout this protocol as an example species. Handling must be done without stress caused by pulling on fur from e.g. latex or nitrile gloves: this can be best achieved using bare hands, in consultation with the facility veterinarian and employee health officials. If this is not possible, soft cotton gloves should be worn over barrier gloves to avoid pulling on the rats' fur.</li>
 <li>Sterile Field Preparation. Prior to beginning surgery, the surgical area is prepared, a sterile field prepared around the stereotaxic apparatus and a sterile drape placed across the apparatus, covering a heating pad that is used to maintain rat body temperature. A circulating warm water pad with accurate thermostat is used to prevent overheating of the animal.</li>
 <li>Anesthesia induction. The isoflurane vaporizer is checked to contain optimal levels of isoflurane and connected to the induction chamber. Connections are checked to confirm a closed loop. The air-handling vacuum system is checked to be operational. Isoflurane vaporization is switched on and the animal placed into the induction chamber using a 5% isoflurane in oxygen mix: that is, 5% isoflurane added into a stream of 100% oxygen and delivered to the induction chamber.</li>
 <li>Placement into apparatus. Animals are secured in the stereotaxic apparatus using earbars and a tooth bar. Correct earbar placement results in an immobile head and ears that lie flat along the earbars. Animals are rapidly secured and the nose inserted into a purpose-designed anesthetic nosecone; delivery of vaporized isoflurane is switched from the induction chamber to the nosecone and the anesthetic mix adjusted to deliver 2-3% isoflurane into the oxygen stream going to the nosecone.</li>
 <li>Confirmation of anesthesia. A surgical anesthetic plane is confirmed by delivering a hard pinch to the foot and a puff of air to the eye; neither should cause any response. These tests are repeated at roughly 15 min intervals throughout surgery to confirm maintenance of a surgical anesthetic plane. Hair is removed from the incision site prior to entering the sterile field.</li>
</ol>
2. Surgery
<ol>
 <li>Animal initial treatment. Ophthalmic ointment is applied to each eyeball to prevent drying. 1 ml sterile saline is given s.c. to prevent any dehydration during surgery, and the body is covered with a sterile drape. Betadine is applied to the scalp, and swabbed from the center out with a cotton swab; 70% ethanol is swabbed similarly, and the two swabbing steps are repeated twice more to ensure an appropriate incision site. Skin contact time for betadine and alcohol should be at least 3 min prior to incision. Anesthesia is maintained and checked regularly throughout surgery; a single s.c. injection of carprofen 5 g/kg) is given to initiate analgesia, and aseptic technique is used. Analgesia is given following induction of anesthesia to minimize any stress from injection.</li>
 <li>Incision and skull preparation. A 3-4 cm cut is made sagitally in the center of the skull. A 1:1 mix of bupivicaine:epinephrine is applied topically to provide further analgesia and minimize bleeding. The scalp is held away from the incision using surgical clips and sterile swabs are used to remove overlying membranes from the skull. Coordinates for drilling are measured using bregma as a reference point, marked using a sterile drill bit (or, alternatively, a hand-held cautery device), and re-confirmed for accuracy before drilling starts. Coordinates for specific brain regions of interest (e.g. here the hippocampus) are determined using a brain atlas. For hippocampal microdialysis, we use a drill site at 5.6 mm posterior to bregma, +5.0 lateral, and 3.0 ventral from dura.</li>
 <li>Drilling. Three holes are drilled through the skull, with care being taken to apply only minimal force such that trauma to the dura and underlying membranes is minimal or absent. One hole is at the measured site of cannula insertion; the other two are positioned as convenient for insertion of skull screws. Purpose-sized screws (e.g. 1.17 mm self-tapping screws; Fine Science Tools) are inserted into these holes without impacting the brain beneath, and are used as anchor points for subsequent dental cement application.</li>
 <li>Cannula placement and closure. The cannula is positioned at insertion coordinates, which are re-confirmed to be the site of the hole drilled, and is then slowly lowered to the target depth. Once correctly positioned, the cannula is secured in place with dental cement. If needed, a single sterile surgical suture is used to close the wound. A stylet is inserted into the cannula to maintain patency. In this protocol we use a CMA12 probe and guide cannula (CMA/Microdialysis).</li>
 <li>Acute post-surgical care. 3 ml sterile saline is given s.c. to continue hydration; a single s.c. injection of carprofen 5 g/kg) is also given to initiate analgesia. Animals are removed from the apparatus and placed in a warmed recovery room, in a clean cage, and monitored until they are fully recovered from anesthesia. Full recovery is assessed by restoration of righting reflex and normal locomotion. Animals are then returned to their home cage and regular holding room.</li>
 <li>Short-term follow-up care. Cages of animals that have undergone surgery are marked with the date of surgery. Animals are monitored at least once each day for at least three days following surgery, and given one carprofen chewable tablet (2 mg) on the day after surgery and each of the two following days. If animals do not consume the carprofen, injectable carprofen may be given to ensure adequate analgesia. Monitor both the overall health of the animal and the state of the wound site (for infection, redness, etc.) following institutional animal care guidelines and seeking assistance or advice from animal care staff or the veterinarian if needed. Importantly, note that appropriate handling and acclimation to the experimenter is essential: animals should be handled extensively, including manipulation of the cannula, until no sign of nervousness or stress remains when handled by the experimenter.</li>
 <li>Subsequent animal care and treatment. Animals will follow appropriate testing and euthanasia procedures according to the approved protocol and their specific experimental group.</li>
</ol>
3. Microdialysis (mD)
<ol>
 <li>Perfusate preparation. Artificial extracellular fluid (aECF) is made with 153.5 mM Na, 4.3 mM K, 0.41 mM Mg, 0.71 mM Ca, 139.4 mM Cl, 1.25 mM glucose, buffered at pH 7.49. NOTE that accurate fluid composition is essential: inaccuracies or use of other, less physiological fluids such as Ringer's or PBS for microdialysis will result in markedly erroneous results9. Specifically, note that the ionic composition of the extracellular fluid (ECF) is NOT the same as that of the CSF, as we showed in a 2004 detailed study of the hippocampal ECF9. On the day of testing, bovine serum albumin (BSA) should be added at 2% weight/weight and fully dissolved; this reduces loss of peptides such as insulin from adherence to the tubing, and also reduces the risk of fluid loss (ultrafiltration) at the probe membrane. After preparation, the perfusate should be filtered through a .2 &mu;m filter.</li>
 <li>If a treatment such as insulin is to be delivered to the brain region of interest (here the hippocampus), prepare this treatment using an aliquot of the prepared aECF with the appropriate drug concentration. For insulin, a concentration of 400 nM (66.7 &mu;U/&mu;l) for delivery via reverse microdialysis has been shown to affect hippocampal metabolic and cognitive function10. Note that the resulting tissue concentration of insulin is not measured here and remains unknown.</li>
 <li>Setting up mD probe and lines. Prepare a fresh microdialysis probe and line the day before testing. Create two separate lines for "inflow" and "outflow". Use PE50 tubing to connect two 1 meter long pieces of FEP tubing and ensure that there is minimal dead space between the lines. Connect the inflow tubing to a 1 ml Hamilton syringe filled with sterile-filtered, deionized H20 (dH20), and then attach to the probe.</li>
 <li>Microdialysis swivel. To allow for free animal movement while taking measurements, connect a liquid swivel to the inflow and outflow lines, close to the pump, using additional FEP tubing. Remember to take the internal volume of this swivel and tubing into account in considering timing of sample collection (below).</li>
 <li>Setting up mD pump. Turn on the mD pump and run at 1.5 &mu;l/min until you see dH20 exiting the outflow tube on the probe. Then, while the pump is turned off, connect the other line between the outflow tube and a sample collection tube. Run 5 ml through the tube and place the probe in a vial containing sterile dH20 overnight; ensure that the probe tip always remains wet.</li>
 <li>Pre-probing test subject. 24 hr before testing, remove the dummy stylet from the rat's head and insert an mD probe (used only for this purpose, not for sampling) for 10 min. Then replace the dummy stylet and put the rat back into its cage. This procedure is designed to minimize any effect of reactive gliosis on the day of testing11-12 and in our hands, this gives good results that match data from other techniques and appear to reflect hippocampal activation4-5,8,10,13; others have used a similar approach that leaves the probe in place for 24 hr prior to measurement, which is also a good approach if damage to the probe overnight can be avoided.</li>
 <li>Probe equilibration. On the day of microdialysis, fill the Hamilton syringe and scintillation vial with filtered aECF and pump through to equilibrate for 1 hr. If needed, fill a second Hamilton syringe with prepared treatment (for example, insulin-aECF) and place into the syringe pump.</li>
 <li>Probe insertion. When equilibration is complete, remove the dummy stylet and gently insert the equilibrated mD probe into the rat's brain via the cannula. Place the rat into a clear plastic box that contains some of the rat's home bedding. Make sure to counterbalance the tubing: attach a 1.6 ml microcentrifuge tube filled with water to the tubing outside the cage in such a way that gravity keeps the microdialysis lines unkinked but not taut. Let probe equilibrate in rat for 2 hr. At the start of this period, confirm that flow of perfusate is unblocked: this is most easily done by collecting perfusate outflow over a defined period and weighing the sample to confirm that the expected volume is exiting the system. Any probe that yields &lt;90% of expected volume, and does not self-correct after removal and re-insertion, should be replaced (otherwise edema at the probe tip will result). If flow is persistently low or absent, check all connections for leakage; failing that, disconnect tubing in stages to see whether the blockage can be isolated. If the problem is not identified, replace the probe; if the problem does not resolve, it may be necessary to start afresh with new probe and lines from Section 3.3. The two-hour equilibration period permits the blood-brain barrier to reseal around the probe and avoid any acute effect of probe insertion.</li>
 <li>Collecting samples. Ensure that the correlation between sample dialysis (from brain) and collection (into tube) is accurately calculated. For instance, using two meters of FEP tubing between swivel and probe, there is a total volume between probe and collection of 30 &mu;l and hence it takes 20 min at 1.5 &mu;l/min flow rate for sample to pass through the probe, tubing, connectors and swivel to reach the end of tubing for collection. When collecting samples ensure that you collect enough volume to have concentration necessary for your assays. Here, we will use 5 min sample bins, and thus collect 7.5 &mu;l of dialysate in each collection tube.</li>
 <li>Baseline samples. Once equilibration is complete, begin sample collection. Collect at least three samples while the rat is at rest in the home chamber to establish a stable baseline of measurements.</li>
 <li>Treatment timing. Take care to calculate required timing for initiation of treatment prior to the experiment: remember that just as there is a time lag between dialysis and sample collection, there is a lag (usually identical) between perfusate leaving the syringe and arriving at the animal's hippocampus. Hence, in our setup with a 30 &mu;l volume between syringe and probe, change the syringe to that containing treatment-perfusate 20 min before you wish the treatment to begin arriving at the hippocampus.</li>
 <li>Changing syringes. A liquid switch can be used if desired but is not necessary: at the appropriate time, simply disconnect the inflow line from the control-perfusate syringe and rapidly connect to the treatment-perfusate syringe. This should take no more than 5 sec to avoid significant interruption to the perfusate flow. NOTE that this process should be reversed after the desired dosage has been delivered, if a time-limited delivery of treatment is desired. Alternatively, treatment may continue for the duration of sampling (see Figure 2). Air bubbles should not be introduced into the line during switching of syringes, as they may accumulate at the dialysis membrane and reduce probe efficiency.</li>
 <li>Behavioral testing. If performing a behavioral task, follow that procedure (see section 4 below). NOTE that coordination with delivery of treatment to the hippocampus is important. For instance, delivery of insulin should be timed to occur 10 min prior to beginning testing 14. Hence, the switch to an insulin-containing perfusate should occur 30 min prior to behavioral testing (20 min for perfusate to pass through the lines plus 10 min desired delivery time ahead of testing).</li>
 <li>Completion of sample collection. After collecting the desired samples, gently remove the probe from the animal's head, again remembering to take into account time lag between dialysis and sample collection. Return the animal to its home cage and observe closely for any post-experimental change in health or behavior until the animal is killed and the brain removed to confirm correct probe placement. If sacrifice will not occur immediately, return the dummy stylet to the cannula to avoid any introduction of foreign material.</li>
 <li>Analysis. The analytical methodology will vary depending on the analyte(s) of interest. Because dialysis sampling does not allow for complete analyte equilibration at the probe membrane, sample concentrations should be corrected to give ECF concentrations using the zero-net-flux method15,16.</li>
</ol>
4. Behavioral Testing
<ol>
 <li>Placement on maze. After baseline samples (i.e., after at least three samples have completed dialysis, but may still be in the outflow tubing being collected), gently move the rat into the behavioral testing apparatus. Any appropriate task may be used; the data in Figure 1 were collected using a four-arm plus-shaped maze and measuring spontaneous alternation, which is a measure of spatial working memory: the rat is initially placed in the center of the maze and allowed to explore freely13,17-18. Because any behavioral test may be used, the focus of this protocol is not on the specific task used as an example here (which is detailed elsewhere13,19,20) but in brief, animals are allowed to explore the maze (the period of the grey box in Figure 1) and use hippocampally-dependent processes to retain memory of which arms have recently been visited.</li>
 <li>Dialysis tubing movement during testing. Hold the tubing such that it moves freely, neither hindering the rat's movement nor being permitted to move in front of the animal and distract it. Stay in place and minimize your own movement so that you do not affect the rat's behavior.</li>
 <li>Continued sample collection. As during baseline, move the outflow tubing to a new collection tube every 5 min.</li>
 <li>Alternation testing. Allow the animal to freely explore the maze for 20 min. Record the sequence and timing of arm entries either using a video recording or by hand for later task performance analysis 13,20.</li>
</ol>
5. Post-testing
<ol>
 <li>Final sample collection. Following testing, remove rat gently from maze and return to control chamber. Continue microdialysis sample collection for at least four samples to cover the period of recovery from task performance.</li>
 <li>Probe removal. After all samples have been collected, gently remove the probe from the animal's head and place into a storage vial; return the animal to its home cage.</li>
 <li>Probe storage. After returning animal to the home cage and completing collection of any remaining dialysate, rinse the probe thoroughly with dH20 and store in a scintillation vial filled with dH20 and covered with parafilm. Flush the tubing by switching to a syringe containing a solution of 1:10,000 Kathon in dH20 to prevent microbe growth. Probes can be re-used as long as they maintain good flow and have no damage to the membrane; with care this should routinely be in excess of ten uses.</li>
 <li>Histology. Kill the animal and remove the brain. Slice on a cryostat and use standard histological techniques (e.g. cresyl violet staining) to confirm correct probe placement.</li>
</ol>

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No conflicts of interest declared.

All solutions used in microdialysis should be filtered immediately prior to use, using a .2 &mu;m filter. After insertion of the probe into the guide cannula, observe to confirm that flow is unimpeded and sample collection is occurring. If flow stops after insertion, the most likely cause is damage to the probe membrane caused by insufficient care on insertion, and a fresh probe must be substituted.
As noted above, a key advantage to these methods is the lack of confound, permitting both beha...

<table border="1">
 <tbody>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>The majority of reagents are standard laboratory grade and can be obtained from a supplier of choice. Similarly, equipment such as syringe pumps and tubing can be used from any of several manufacturers. Specific items used here for which details are important include:</td>
 </tr>
 <tr>
 <td>CMA 12 microdialysis probes</td>
 <td>CMA/ Microdialysis</td>
 <td>CMA-12-XXX</td>
 <td>These are available in various membrane lengths and cutoffs, indicated by specific codes in the 'XXX.'</td>
 </tr>
 <tr>
 <td>Human insulin (Humulin)</td>
 <td>Eli Lilly</td>
 <td>N/a</td>
 <td></td>
 </tr>
 <tr>
 <td>Liquid swivel</td>
 <td>Instech</td>
 <td>375/D/22QM</td>
 <td>This specific swivel has very low torque and internal volume, as well as a nonreactive quartz lining.</td>
 </tr>
 </tbody>
 </table>

hippocampal insulin microinjection and in vivo microdialysis during spatial memory testing

Glucose metabolism is a useful marker for local neural activity, forming the basis of methods such as 2-deoxyglucose and functional magnetic resonance imaging. However, use of such methods in animal models requires anesthesia and hence both alters the brain state and prevents behavioral measures. An alternative method is the use of in vivo microdialysis to take continuous measurement of brain extracellular fluid concentrations of glucose, lactate, and related metabolites in awake, unrestrained animals. This technique is especially useful when combined with tasks designed to rely on specific brain regions and/or acute pharmacological manipulation; for example, hippocampal measurements during a spatial working memory task (spontaneous alternation) show a dip in extracellular glucose and rise in lactate that are suggestive of enhanced glycolysis1-3,4-5, and intrahippocampal insulin administration both improves memory and increases hippocampal glycolysis6. Substances such as insulin can be delivered to the hippocampus via the same microdialysis probe used to measure metabolites. The use of spontaneous alternation as a measure of hippocampal function is designed to avoid any confound from stressful motivators (e.g. footshock), restraint, or rewards (e.g. food), all of which can alter both task performance and metabolism; this task also provides a measure of motor activity that permits control for nonspecific effects of treatment. Combined, these methods permit direct measurement of the neurochemical and metabolic variables regulating behavior.

Rats should recover rapidly from surgery and be alert, motile and active within 30 min following cessation of anesthesia. The impact of the cannula cap should be minimal if the surgery is performed cleanly and minimal dental cement is used. If any sign of infection is noticed during the post-operative monitoring, or the rat is in any way showing signs of distress or discomfort, immediately terminate the experiment; this should be extremely rare. During the handling prior to testing, animals should be alert, freely motile...

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Hippocampal Insulin Microinjection and In vivo Microdialysis During Spatial Memory Testing

Modulation of hippocampally-dependent spatial working memory by direct intrahippocampal microinjection, accompanied and followed by in vivo microdialysis for metabolites in conscious, behaving animals.

Hippocampal Insulin Microinjection and In vivo Microdialysis During Spatial Memory Testing

Glucose metabolism is a useful marker for local neural activity, forming the basis of methods such as 2-deoxyglucose and functional magnetic resonance ...

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Research

JoVE Journal

Neuroscience

16.4K Views.  University at Albany. Modulation of hippocampally-dependent spatial working memory by direct intrahippocampal microinjection, accompanied and followed by in vivo microdialysis for metabolites in conscious, behaving animals.

Video: Hippocampal Insulin Microinjection and In vivo Microdialysis During Spatial Memory Testing

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Microdialysis of Ethanol During Operant Ethanol Self-administration and Ethanol Determination by Gas Chromatography

Operant self-administration methods are commonly used to study the behavioral and pharmacological effects of many drugs of abuse, including ethanol. However, ethanol is typically self-administered orally, rather than intravenously like many other drugs of abuse. The pharmacokinetics of orally administered drugs are more complex than intravenously administered drugs. Because understanding the relationship between the pharmacological and behavioral effects of ethanol requires knowledge of the time course of ethanol reaching the brain during and after drinking, we use in vivo microdialysis and gas chromatography with flame ionization detection to monitor brain dialysate ethanol concentrations over time. 
Combined microdialysis-behavioral experiments involve the use of several techniques. In this article, stereotaxic surgery, behavioral training and microdialysis, which can be adapted to test a multitude of self-administration and neurochemical centered hypotheses, are included only to illustrate how they relate to the subsequent phases of sample collection and dialysate ethanol analysis. Dialysate ethanol concentration analysis via gas chromatography with flame-ionization detection, which is specific to ethanol studies, is described in detail. Data produced by these methods reveal the pattern of ethanol reaching the brain during the self-administration procedure, and when paired with neurochemical analysis of the same dialysate samples, allows conclusions to be made regarding the pharmacological and behavioral effects of ethanol.

A method to determine the time course of ethanol concentration in the brains of rats during operant ethanol self-administration is described. Gas chromatography with flame ionization detection is used to quantify ethanol in the dialysate samples, because it has the sensitivity required for the small volumes that are generated.

Operant self-administration methods are commonly used to study the behavioral and pharmacological effects of many drugs of abuse, including ethanol.  ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

High-resolution In Vivo Manual Segmentation Protocol for Human Hippocampal Subfields Using 3T Magnetic Resonance Imaging

The human hippocampus has been broadly studied in the context of memory and normal brain function and its role in different neuropsychiatric disorders has been heavily studied. While many imaging studies treat the hippocampus as a single unitary neuroanatomical structure, it is, in fact, composed of several subfields that have a complex three-dimensional geometry. As such, it is known that these subfields perform specialized functions and are differentially affected through the course of different disease states. Magnetic resonance (MR) imaging can be used as a powerful tool to interrogate the morphology of the hippocampus and its subfields. Many groups use advanced imaging software and hardware (&#62;3T) to image the subfields; however this type of technology may not be readily available in most research and clinical imaging centers. To address this need, this manuscript provides a detailed step-by-step protocol for segmenting the full anterior-posterior length of the hippocampus and its subfields: cornu ammonis (CA) 1, CA2/CA3, CA4/dentate gyrus (DG), strata radiatum/lacunosum/moleculare (SR/SL/SM), and subiculum. This protocol has been applied to five subjects (3F, 2M; age 29-57, avg. 37). Protocol reliability is assessed by resegmenting either the right or left hippocampus of each subject and computing the overlap using the Dice&#39;s kappa metric. Mean Dice&#39;s kappa (range) across the five subjects are: whole hippocampus, 0.91 (0.90-0.92); CA1, 0.78 (0.77-0.79); CA2/CA3, 0.64 (0.56-0.73); CA4/dentate gyrus, 0.83 (0.81-0.85); strata radiatum/lacunosum/moleculare, 0.71 (0.68-0.73); and subiculum 0.75 (0.72-0.78). The segmentation protocol presented here provides other laboratories with a reliable method to study the hippocampus and hippocampal subfields in vivo using commonly available MR tools.

High-resolution In Vivo Manual Segmentation Protocol for Human Hippocampal Subfields Using 3T Magnetic Resonance Imaging

The goal of this manuscript is to study the hippocampus and hippocampal subfields using MRI. The manuscript describes a protocol for segmenting the hippocampus and five hippocampal substructures: cornu ammonis (CA) 1, CA2/CA3, CA4/dentate gyrus, strata radiatum/lacunosum/moleculare, and subiculum.

The human hippocampus has been broadly studied in the context of memory and normal brain function and its role in different neuropsychiatric disorders has ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Immunohistochemical Visualization of Hippocampal Neuron Activity After Spatial Learning in a Mouse Model of Neurodevelopmental Disorders

Induction of phosphorylated extracellular-regulated kinase (pERK) is a reliable molecular readout of learning-dependent neuronal activation. Here, we describe a pERK immunohistochemistry protocol to study the profile of hippocampal neuron activation following exposure to a spatial learning task in a mouse model characterized by cognitive deficits of neurodevelopmental origin. Specifically, we used pERK immunostaining to study neuronal activation following Morris water maze (MWM, a classical hippocampal-dependent learning task) in Engrailed-2 knockout (En2-/-) mice, a model of autism spectrum disorders (ASD). As compared to wild-type (WT) controls, En2-/- mice showed significant spatial learning deficits in the MWM. After MWM, significant differences in the number of pERK-positive neurons were detected in specific hippocampal subfields of En2-/- mice, as compared to WT animals. Thus, our protocol can robustly detect differences in pERK-positive neurons associated to hippocampal-dependent learning impairment in a mouse model of ASD. More generally, our protocol can be applied to investigate the profile of hippocampal neuron activation in both genetic or pharmacological mouse models characterized by cognitive deficits.

We describe an immunohistochemistry protocol to study the profile of hippocampal neuron activation following exposure to a spatial learning task in a mouse model characterized by cognitive deficits of neurodevelopmental origin. This protocol can be applied to both genetic or pharmacological mouse models characterized by cognitive deficits.

Induction of phosphorylated extracellular-regulated kinase (pERK) is a reliable molecular readout of learning-dependent neuronal activation. Here, we ...

Assessing Spatial Learning and Memory in Small Squamate Reptiles

Clinical research has leveraged a variety of paradigms to assess cognitive decline, commonly targeting spatial learning and memory abilities. However, interest in the cognitive processes of nonmodel species, typically within an ecological context, has also become an emerging field of study. In particular, interest in the cognitive processes in reptiles is growing although experimental studies on reptilian cognition are sparse. The few reptilian studies that have experimentally tested for spatial learning and memory have used rodent paradigms modified for use in reptiles. However, ecologically important aspects of the physiology and behavior of this taxonomic group must be taken into account when testing for spatially based cognition. Here, we describe modifications of the dry land Barnes maze and associated testing protocol that can improve performance when probing for spatial learning and memory ability in small squamate reptiles. The described paradigm and procedures were successfully used with male side-blotched lizards (Uta stansburiana), demonstrating that spatial learning and memory can be assessed in this taxonomic group with an ecologically relevant apparatus and protocol.

This paper describes a modification of the Barnes maze, a standard rodent paradigm used to assess spatial memory and learning, for use in small squamate reptiles.

Clinical research has leveraged a variety of paradigms to assess cognitive decline, commonly targeting spatial learning and memory abilities. However, ...

An In Ovo Model for Testing Insulin-mimetic Compounds

Elevated blood glucose levels in type 2 diabetes mellitus (T2DM), a complex and multifactorial metabolic disease, are caused by insulin resistance and &#946;-cell failure. Various strategies, including the injection of insulin or the usage of insulin-sensitizing drugs, were pursued to treat T2DM or at least reduce the symptoms. In addition, the application of herbal compounds has attracted increasing attention. Thus, it is necessary to find efficient test systems to identify and characterize insulin-mimetic compounds. Here we developed a modified chick embryo model, which enables testing of synthetic compounds and herbal extracts with insulin-mimetic properties. Using a fluorescence microscopy-based primary screen, which quantifies the translocation of Glucose transporter 4 (Glut4) to the plasma membrane, we were able to identify compounds, mainly herbal extracts, which lead to an increase of intracellular glucose concentrations in adipocytes. However, the efficacy of these substances requires further verification in a living organism. Thus, we used an in-ovo approach to identify their blood glucose-reducing properties. The approval by an ethics committee is not needed since the use of chicken embryos during the first two-thirds of embryonic development is not considered an animal experiment. Here, the application of this model is described in detail.

An In Ovo Model for Testing Insulin-mimetic Compounds

In this study, we describe an in-ovo system as a promising tool to test insulin-mimetic compounds in a living organism. The main advantages include adequate throughput rates and acceptable costs, which enable the identification of phytochemicals with insulin-mimetic properties.

Elevated blood glucose levels in type 2 diabetes mellitus (T2DM), a complex and multifactorial metabolic disease, are caused by insulin resistance and ...