We thank Bernard Foucart for technical assistance. This work was supported by the Belgian Fund for Scientific Research (F.R.S.-FNRS) and by the Queen Elisabeth Fund for Medical Research. Agn&egrave;s Villers is Research Fellow at the Belgian Fund for Scientific Research.

All animal procedures were carried out in accordance with National Institutes of Health regulations for the care and use of animals in research and with the agreement of the local ethics committee.
1. Preparation of Artificial Cerebro-spinal Fluid
The same media is used to dissect, cut and perfuse slices (1 ml/min) during the resting period and the electrophysiological recordings. This media is composed of 124 mM NaCl, 4.4 mM KCl, 26 mM NaHCO3, 1 mM NaH2PO4, 2.5 mM CaCl2, 1.3 mM MgSO4 and 10 mM D-glucose.
<ol>
	<li>Weigh the different components for 2 L of ACSF except MgSO4 which is already in solution (1 M). Using a standard solution allows to be sure of the exact concentration of Mg2+ because MgSO4 in powder is highly hygroscopic. Ca2+:Mg2+ ratio greatly influences LTP induction and maintenance 14 and thus their proportions must always be respected.</li>
	<li>Put all the components except CaCl2 in a glass graduated cylinder and bring to 2 L with distilled H2O.</li>
	<li>Stir vigorously. When everything is dissolved, add carbogen through an aquarium bubbler and tubing attached to the tank (95% O2, 5% CO2). Wait a few minutes and add calcium chloride when pH is fixed at 7.4 by the action of bicarbonate buffer.</li>
	<li>Keep 600 ml in a refrigerated rectangular glass dish and cool down to 4 &deg;C with ice around the dish. This media will be used for hippocampus dissection.</li>
	<li>Filter the remaining 1,400 ml and keep in an Erlenmeyer flask.</li>
</ol>
2. Preparation of the Electrophysiological Rig and the Brain Slice Recording Chamber
The perfusion circuit is made of 2 peristaltic pumps, one pumping fresh fluid from a reserve tank and the other pumping used fluid from the recording chamber to a bin. Fresh fluid is added to a home-made perfusion system where it is re-oxygenated and warmed before reaching the interface recording chamber by gravity (Figure 1A). The interface recording chamber was designed by FST (Fine Science Tools, Vancouver, Canada, Figure 1B). Tissue slices are placed on woven net filter attached to a removable well ring and can be continuously maintained under interface conditions by adjusting the height of the suction well needle. A home-made plastic tube blocked at its end and perforated laterally (0.5 mm) is fixed to the suction needle to increase level stability in the chamber. The recording head is mounted over a water bath containing a gas dispersion stone which helps maintain a humid environment by passing moist air through deflection ports in the recording head (to decrease water droplet formation). Bath and medium temperature are controlled by continuously varying the level of DC current through a silicone rubber-embedded heater element in the water bath. Thermistors in the water bath and recording wells allow the chamber temperature to be set and precisely monitored at these sites.
The heating system of the chamber is completed by a global heating system controlling the temperature of the entire rig. The software developed by researchers at the University of Edinburgh (<a href="http://www.etcsystem.com" target="_blank">www.etcsystem.com</a>) monitors and equalizes the temperature of the oxygenated air, the brain slice, the electrodes and the remaining rig providing a stable environment for long-term recordings (Figure 1C). The temperature used for mouse hippocampal slices is 28 &deg;C.
<ol>
	<li>Rinse all the perfusion circuit with distilled H2O for a minimum of 20 min and start the heating systems.</li>
	<li>Start the carbogen bubbling in the circuit. Carbogen arrives in the home-made perfusing tubes through filter candles and in the water bath below the recording chamber. The flow rate in the water bath is controlled by a flow meter. If the flow rate is too slow, tissue could become hypoxic. Conversely, if the flow rate is too high, the gas might not be sufficiently warmed and moisturized leading to drying of the tissue. High gas flow rate could also increase the chance of water spraying over the tissue from the water bath below leading to osmotic shock. This can be prevented by adding bits of nylon mesh (100 &mu;m) at the outlet of the deflection ports.</li>
	<li>Drain the circuit and fill up with filtered ACSF.</li>
	<li>Put the ring which will support the slice in the holding chamber. A woven net filter with mesh openings ranging from 500 to 750 &mu;m is stretched and attached to the ring with two-part resin epoxy glue. The woven net filter is made of 87% polyamide and 13% elastane (panty 15 den). Glue must be applied carefully to avoid the formation of reliefs at the surface of the ring.</li>
	<li>Carefully remove all the air bubbles from the circuit.</li>
	<li>Adjust the level of ACSF in the recording chamber with the screw of the suction needle. The speed of the peristaltic pumps is adjusted to 1 ml/min for the inlet pump and to 5 ml/min for the outlet pump.</li>
</ol>
The perfusion system is placed on a rigid vibration-resistant table and surrounded by a Faraday cage.
3. Preparation of the Dissection Area
Surgical instruments: a scalpel with a #11 blade, small standard dissecting scissors, spring scissors, curved forceps, two Heidemann spatulae and a spoon.
Supplementary material: a guillotine, an underpad, a McIlwain tissue chopper with a razor blade, filter paper, a glass Pasteur pipette with a rubber teat, 2 plastic Pasteur pipettes one of which with a wide mouth, a Petri dish, a metallic cylinder of 7 mm diameter (Figure 2A).
<ol>
	<li>In the dissecting dish filled with cold ACSF (prepared in Step 1), immerse a refrigerated glass lid from a staining dish, wrapped in filter paper. Carefully remove air bubbles and oxygenate the ACSF through an aquarium bubbler (Figure 2A).</li>
	<li>Lay out instruments in order of use. Add three layers of filter paper on the tissue chopper plate and a new razor blade cleaned with distilled water and ether (Figure 2B). The razor blade must be horizontal when it touches the paper. Blade force is adjusted to half of maximal value and speed to approximately one third of maximal value.</li>
	<li>Attentively check if everything is ready (instruments, temperature ...) because you will not have time afterwards.</li>
	<li>Ask someone to spray the dissection with a Pasteur pipette filled with cold ACSF.</li>
	<li>Anesthetize the mouse with Nembutal IP (100 mg/kg) before decapitation. Halothane anesthesia can also be used. We have compared both methods and obtained same results. Decapitation must be performed under anesthesia but cervical dislocation can also be used. However cervical dislocation needs a very good practice to avoid animal suffering.</li>
	<li>Dissect on the underpad as quickly as possible under cold ACSF continuous shower.</li>
</ol>
4. Brain Removal
<ol>
	<li>While holding still the head with the index finger and the thumb of one hand on the muzzle, make an incision with the dissecting scissors along the middle of the top of the head starting at the edge of the guillotine cut and running rostrally to the frontal bone.</li>
	<li>Cut through the cutaneous muscle on each side of the head to fully expose the skull plates and remove the muscles at the caudal side of the head.</li>
	<li>With the dissecting scissors, cut the temporalis muscle on each side, along the temporalis plate, then cut the frontal plates in the middle, transversally. Then, make a little cut on the occipital bone, between the two plates.</li>
	<li>For each side, cut at the caudal base of the occipital plates.</li>
	<li>Cut along the sagittal suture with spring scissors.</li>
	<li>With forceps, remove the skull halves by spreading them away from each other.</li>
	<li>With the scalpel, cut transversally just after the olfactory bulb and just before the cerebellum then plunge the extracted brain into the dissecting dish with cold ACSF.</li>
</ol>
5. Hippocampal Dissection
<ol>
	<li>Once the brain is immerged in the dissecting dish, sever the two hemispheres from each other with a scalpel inserted in the middle.</li>
	<li>The dissection of hippocampus is performed with spatulae under visual control through a binocular surgical microscope (X25, Figure 2A). On one hemisphere, carefully spread the structures to see the lateral ventricle. Remove the brain stem and diencephalon. This is done by applying the spatulae on the frontal cortex on one side and on the diencephalon on the other side. Take care to not touch the hippocampus with the spatulae and to not stretch it during tissue sectioning.</li>
	<li>Sever the fornix then gently push the hippocampus out of the cortex (roll away), by inserting a spatula in the ventricle.</li>
	<li>When the hippocampus is extracted, remove excess cortex tissue and remaining blood vessels.</li>
</ol>
6. Cutting of the Slices
<ol>
	<li>With a wide mouth plastic Pasteur pipette, transfer the hippocampus in a spoon with its alvear surface upwards (convex side).</li>
	<li>Remove excess fluid with a standard plastic Pasteur pipette.</li>
	<li>Tip up the spoon vertically nearly touching the filter paper of the chopper (Figure 2B) and drop off the hippocampus from the spoon, by rapidly touching the filter paper then move back the spoon.</li>
	<li>Orientate the hippocampus and slice it transversally to 400 &mu;m with the chopper (see Figure 2C for orientation of the hippocampus relative to the razor blade). Act as quickly as possible.</li>
	<li>Remove the filter paper with the sliced hippocampus and wrap it around the metallic cylinder to spread the slices a little. Then, free the slices with sprays of ACSF using a standard plastic Pasteur pipette and collect them in a Petri dish filled with cold ACSF.</li>
</ol>
7. Incubation of Slices in Interface
<ol>
	<li>Select slices with a standard plastic Pasteur pipette and rapidly place them in the recording chamber. Slices recover from the dissection trauma directly in the recording chamber, in interface at 28 &deg;C.</li>
	<li>The slice will most likely sink. Lower the fluid level to reach the slice level, and then raise it to make it float.</li>
	<li>Turn the slice in the correct position while lowering the level. Slices are always orientated in the same way to facilitate the positioning of electrodes (2 stimulating and one recording electrodes) in the CA1 region.</li>
	<li>Stop when the fluid is in interface. A "meniscus" of media around the slice is indicative of a sufficient level of media. The net filter must be saturated with media but not completely submerged.</li>
	<li>Keep the chamber covered with filter papers placed over the perforated lid.</li>
	<li>Let the slice rest for at least 1.5 hr at 28 &deg;C.</li>
</ol>
8. Recording of Synaptic Responses
<ol>
	<li>Recording electrode: glass capillaries are pulled to obtain a tip resistance of 2-5 M&Omega; when filled with ACSF. A small piece of filter paper is placed around the tip of the electrode and bone wax is applied at the extremity to reduce condensation. Stimulating electrodes: platinum-iridium bipolar cluster electrodes with a 12.5 &mu;m diameter are purchased from FHC (USA). With proper care, each electrode can be used at least 30 times (Figure 1B).</li>
	<li>Position the electrodes in the stratum radiatum of the CA1 region, all the electrodes lined up (Figure 3A). Electrodes are lowered 75 to 150 &mu;m under the surface of the slice with Narishige micromanipulators. The two stimulating electrodes are placed to stimulate two distinct bundles of Schaffer collaterals. When the electrodes are lowered onto the slice, filter papers are carefully placed all around to close the chamber.</li>
	<li>Biphasic stimulation (0.08 msec pulse duration per half-wave) is performed at constant voltage with a Grass stimulator connected to SIU-V isolation units. Maximal response is checked by increasing stimulus intensity from 2 V to maximum 12 V. Field EPSPs are amplified 1,000 times with a WPI ISO-80 amplifier and filtered at 10 Hz and 10 kHz. The signal is then sent to a PC through a National Instrument A/D converter. Stimulation, data acquisition and analysis are performed using the WinLTP program (<a href="http://www.winltp.com">www.winltp.com</a>). Field EPSPs are recorded at 40% of the maximum amplitude obtained in an input-output curve. For each slice, the fEPSP slopes are normalized against the average slope over the 30 min preceding LTP induction.</li>
	<li>LTP is triggered by applying a single train of stimulation (100 Hz) at test strength on one pathway while the second pathway serves as a control.</li>
</ol>
9. Cleaning of the Setup
<ol>
	<li>Rinse all the circuit with a 3% solution of hydrogen peroxide (H2O2) for at least 10 min, then drain. The circuit will be carefully rinsed with distilled water before starting any experiment. The use of H2O2 is not absolutely required as other labs use only distilled water for cleaning but we have noticed the deposition of dark residues in the tubing system when using only water.</li>
	<li>Replace water in the water bath beneath the recording chamber.</li>
	<li>Bath the stimulating electrodes tips in alcohol for 5 min.</li>
</ol>

No conflicts of interest declared.

We have developed in our laboratory a protocol resulting from the combination of methods developed and used by other laboratories having a big expertise in LTP recordings 11,17. This protocol is adapted to adult mouse hippocampus and can be used in animals of any age and any background genotype. It also allows the analysis of LTP in transgenic mice developing neurodegenerative diseases like Alzheimer's disease 18,19.
Utilization of this protocol for rat hippocampal slice...

Nowadays, there is limited understanding of how complex memories are stored and recalled at the neuronal circuit level. However, a unifying hypothesis of memory storage is available and broadly accepted: memories are stored as changes in the strength of synaptic connections between neurons in the central nervous system. On its own, research on synaptic plasticity has largely benefited from two breakthrough discoveries. (1) In a seminal experiment, Bliss and Lomo 1, using the intact anesthetized rabbit, found that delivery of a brief high-frequency (1 sec, 100 Hz) stimulation to the perforant path of the hippocampus caused a long-lasting (several hours) increase in the related synaptic connections. This fascinating phenomenon was called "Long-Term Potentiation" or LTP by Douglas and Goddard in 1975 2. (2) Later on, it was found that a similar phenomenon could be triggered in brain slices (0.4 mm) artificially maintained alive in vitro. The most widely studied LTP was observed in vitro by delivering one or several tetani to a bundle of axons (the so-called Schaffer collaterals) while recording the resulting field excitatory synaptic potential evoked in the pyramidal neurons of the so-called CA1 region. The mechanisms of LTP induction have largely been revealed. Basically, a Ca2+ influx through the NMDA receptors activates enzymes with two consequences: a phosphorylation of AMPA receptors (which increases their efficiency) and an incorporation of extra AMPA receptors in the postsynaptic membrane 3. By contrast, the mechanisms of the maintenance phase of LTP are largely unknown, notably because it is experimentally much more difficult to maintain a slice healthy for many hours than for 30 to 60 min.
A lot of studies have been dedicated to the understanding of LTP mechanisms and interesting theories have been elaborated over the years 4-11. But until now, the precise molecular mechanisms underlying the stable increase in synaptic strength have not been elucidated. This could be partly due to the difficulty to reproduce previous results in different laboratories using different techniques for the preparation and the maintenance of hippocampal slices. In their methodology paper, Sajikumar et al. 12 stressed the importance of experimental conditions for the preparation of rat hippocampal slices and the recording of stable LTP. In this video we present all the optimization steps developed in our laboratory over the years to be able to record a very stable LTP in mouse hippocampal slices.
This optimization has been made from protocols developed and successfully used by other laboratories which study LTP mechanisms in mice 13 and rats 11. It allows experienced researchers to induce and record a very long lasting LTP in adult mice with a high rate of success. The physiological basis of the induced LTP was carefully checked and demonstrated 14. In this methodology paper, we show that any modifications of experimental conditions, like temperature or oxygenation can have a profound impact on LTP maintenance while the dissection procedure can deeply modify slices excitability. It must also be emphasized that the precise control of all these parameters requires a training of several months for novice students.

<table border="1">
	<tbody>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Reagent/Material</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma - Aldrich</td>
			<td>S7653</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma - Aldrich</td>
			<td>S8875</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma - Aldrich</td>
			<td>P9333</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Sigma - Aldrich</td>
			<td>G7528</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Sigma - Aldrich</td>
			<td>S9638</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>MgSO4 1M</td>
			<td>Sigma - Aldrich</td>
			<td>63126</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma - Aldrich</td>
			<td>C4901</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Carbogen</td>
			<td>Air Liquide (Belgium)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Capillaries</td>
			<td>WPI, Inc. (UK)</td>
			<td>TW150-4</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Stimulating Electrodes</td>
			<td>FHC (USA)</td>
			<td>CE2B30</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Surgical tools</td>
			<td>FST (Germany)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Filter paper 84 g/m2</td>
			<td>Sartorius</td>
			<td>FT-3-105-110</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Mesh</td>
			<td>Lycra</td>
			<td>15 den</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Glue</td>
			<td>UHU</td>
			<td>plus endfest300</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Instrument</td>
		</tr>
		<tr>
			<td>Amplifier</td>
			<td>WPI, Inc. (UK)</td>
			<td>ISO-80</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Interface recording chamber</td>
			<td>FST (Germany)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Peristaltic pumps</td>
			<td>Gilson (USA)</td>
			<td>Minipuls 3</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Temperature controller</td>
			<td>University of Edinburgh</td>
			<td><a href="http://www.etcsystem.com" target="_blank">www.etcsystem.com</a></td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tissue Chopper</td>
			<td>Mcllwain</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Stimulators</td>
			<td>Grass (USA)</td>
			<td>S88X + SIU-V</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Program analysis</td>
			<td>WinLTP</td>
			<td><a href="http://www.winltp.com" target="_blank">www.winltp.com</a></td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Micromanipulators</td>
			<td>Narishige</td>
			<td>MM-3 and MMO-220A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td>Leica Microsystem</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>A/D converter</td>
			<td>National Instruments</td>
			<td>NIPCI-6229 M-series</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

improved preparation and preservation of hippocampal mouse slices for a very stable and reproducible recording of long-term potentiation

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory encoding. LTP elicited in the CA1 region of acute hippocampal slices has been extensively studied. However the molecular mechanisms underlying the maintenance phase of this phenomenon are still poorly understood. This could be partly due to the various experimental conditions used by different laboratories. Indeed, the maintenance phase of LTP is strongly dependent on external parameters like oxygenation, temperature and humidity. It is also dependent on internal parameters like orientation of the slicing plane and slice viability after dissection.
The optimization of all these parameters enables the induction of a very reproducible and very stable long-term potentiation. This methodology offers the possibility to further explore the molecular mechanisms involved in the stable increase in synaptic strength in hippocampal slices. It also highlights the importance of experimental conditions in in vitro investigation of neurophysiological phenomena.

This methodology has been used to analyze the properties of long-lasting long-term potentiation induced in acute hippocampal slices from adult C57Bl/6J mice (JANVIER SAS, France) 14. Surprisingly, improvement of the experimental conditions has led to a new way of looking at LTP. We showed that long-lasting increase in synaptic strength did not require the synthesis of new proteins.
Here, we show that LTP induction depends on slices viability and excitability. When dissection of the ...

Watch this Scientific Journal Video about Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation at JoVE.com

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory ...

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26.6K Views.  University of Mons. This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Video: Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Preparation of Oligomeric &beta;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). &beta;-amyloid (A&beta;), a peptide produced in high amounts in AD, is known to reduce Long-Term Potentiation (LTP), a cellular correlate of learning and memory. Indeed, LTP impairment caused by A&beta; is a useful experimental paradigm for studying synaptic dysfunctions in AD models and for screening drugs capable of mitigating or reverting such synaptic impairments. Studies have shown that A&beta; produces the LTP disruption preferentially via its oligomeric form. Here we provide a detailed protocol for impairing LTP by perfusion of oligomerized synthetic A&beta;1-42 peptide onto acute hippocampal slices. In this video, we outline a step-by-step procedure for the preparation of oligomeric A&beta;1-42. Then, we follow an individual experiment in which LTP is reduced in hippocampal slices exposed to oligomerized A&beta;1-42 compared to slices in a control experiment where no A&beta;1-42 exposure had occurred.

Preparation of Oligomeric &amp;#946;-amyloid1-42 and Induction of Synaptic Plasticity Impairment on Hippocampal Slices

One feature of Alzheimer's Disease is the elevation of A&beta;1-42 peptide. Here we provide a protocol for preparing synthetic A&beta;1-42 oligomers, which impairs hippocampal Long-Term Potentiation, a cellular correlate of memory. This procedure is useful for investigating mechanisms of A&beta;-induced pathology and drug screening.

Impairment of synaptic connections is likely to underlie the subtle amnesic changes occurring at the early stages of Alzheimer s Disease (AD). ...

Preparation of Acute Hippocampal Slices from Rats and Transgenic Mice for the Study of Synaptic Alterations during Aging and Amyloid Pathology

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The hippocampus can be extracted quickly and easily from rats and mice and slices remain viable for hours in oxygenated artificial cerebrospinal fluid. Moreover, basic electrophysisologic techniques are easily applied to the investigation of synaptic function in hippocampal slices and have provided some of the best biomarkers for cognitive impairments. The hippocampal slice is especially popular for the study of synaptic plasticity mechanisms involved in learning and memory. Changes in the induction of long-term potentiation and depression (LTP and LTD) of synaptic efficacy in hippocampal slices (or lack thereof) are frequently used to describe the neurologic phenotype of cognitively-impaired animals and/or to evaluate the mechanism of action of nootropic compounds. This article outlines the procedures we use for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and Alzheimer's disease (AD)1-3. Use of aged rats and AD model mice can present a unique set of challenges to researchers accustomed to using younger rats and/or mice in their research. Aged rats have thicker skulls and tougher connective tissue than younger rats and mice, which can delay brain extraction and/or dissection and consequently negate or exaggerate real age-differences in synaptic function and plasticity. Aging and amyloid pathology may also exacerbate hippocampal damage sustained during the dissection procedure, again complicating any inferences drawn from physiologic assessment. Here, we discuss the steps taken during the dissection procedure to minimize these problems. Examples of synaptic responses acquired in "healthy" and "unhealthy" slices from rats and mice are provided, as well as representative synaptic plasticity experiments. The possible impact of other methodological factors on synaptic function in these animal models (e.g. recording solution components, stimulation parameters) are also discussed. While the focus of this article is on the use of aged rats and transgenic mice, novices to slice physiology should find enough detail here to get started on their own studies, using a variety of rodent models.

This article outlines procedures for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and age-related neurodegenerative diseases, such as Alzheimer&#x2019;s disease.

The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

Preparation of an Awake Mouse for Recording Neural Responses and Injecting Tracers

It is well known that anesthesia alters neural response properties in various regions of the brain.13. In the auditory system, fundamental response properties of brainstem neurons including threshold, frequency specificity, and inhibitory sidebands are altered in significant ways under anesthesia1-2. These observations prompted physiologists to seek ways to record from single neurons without the contaminating effects of anesthesia. One result was a decerebrate preparation, where the brainstem was completely transected at the level of the midbrain4. The drawbacks of this preparation are a formidable surgery, the elimination of descending projections from the forebrain, and an inability to use sensory stimulation to examine structures above the midbrain. A different strategy has been to implant electrode arrays chronically to record from single neurons and multiunit clusters while the animal is awake and/or behaving5,6. These techniques however are not compatible with injecting tracer dyes after first electrophysiologically characterizing a brain structure. To avoid altering neural response properties with anesthetics while recording electrophysiological response properties from single neurons, we have adapted a head restraint technique long used in bats7-9 to mouse10-12. Using this method, we are able to conduct electrophysiological recordings over several days in the unanesthetized mouse. At the end of the recording sessions, we can then inject a dye to reconstruct electrode positions and recording sites or inject a tracer so that pathways to and from the recording loci can be determined. This method allows for well isolated single neuron recordings over multiple days without the use anesthetics.

Electrophysiological characterization of neuronal responses is important for understanding brain function and for guiding the placement of dyes for pathway tracing. However, many studies are performed in anesthetized animals. To understand brain function without anesthetics, we developed a method to record neuronal response properties and inject dyes in awake mouse.

It is well known that anesthesia alters neural response properties in various regions of the brain.13.  In the auditory system, fundamental response ...

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

A Simplified Method for Ultra-Low Density, Long-Term Primary Hippocampal Neuron Culture

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic hippocampal neurons can often grow successfully on glass coverslips at high density under serum-free conditions, but low density cultures typically require a supply of trophic factors by co-culturing them with a glia feeder layer, preparation of which can be time-consuming and laborious. In addition, the presence of glia may confound interpretation of results and preclude studies on neuron-specific mechanisms. Here, a simplified method is presented for ultra-low density (~2,000 neurons/cm2), long-term (&#62;3 months) primary hippocampal neuron culture that is under serum free conditions and without glia cell support. Low density neurons are grown on poly-D-lysine coated coverslips, and flipped on high density neurons grown in a 24-well plate. Instead of using paraffin dots to create a space between the two neuronal layers, the experimenters can simply etch the plastic bottom of the well, on which the high density neurons reside, to create a microspace conducive to low density neuron growth. The co-culture can be easily maintained for &#62;3 months without significant loss of low density neurons, thus facilitating the morphological and physiological study of these neurons. To illustrate this successful culture condition, data are provided to show profuse synapse formation in low density cells after prolonged culture. This co-culture system also facilitates the survival of sparse individual neurons grown in islands of poly-D-lysine substrates and thus the formation of autaptic connections.

Low density cultures of primary hippocampal neurons usually require glia feeder layer to supply neurotrophic factors and sustain longevity. We describe here a simplified method to culture ultra-low density neurons on glass coverslips in the presence of a high density neuronal feeder layer, which facilitates investigation of specific neuronal-autonomous mechanisms.

Culturing primary hippocampal neurons in vitro facilitates mechanistic interrogation of many aspects of neuronal development. Dissociated embryonic ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...