This video article is sponsored by Cerebos Pacific Limited. This work is supported by National Medical Research Council Collaborative Research Grant (NMRC-CBRG-0041/2013) and Ministry of Education Academic Research Funding (MOE AcRF- Tier 1 - T1-2012 Oct -02).

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of National University of Singapore.
1. Preparation of Artificial Cerebrospinal Fluid (ACSF)
<ol>
	<li>Prepare the ACSF consisting of (in mM) 124 NaCl, 3.7 KCl, 1.0 MgSO4.7H2O, 2.5 CaCl2.2H2O, 1.2 KH2PO4, 24.6 NaHCO3, and 10 D-glucose. Ensure the pH of the ACSF is between 7.2-7.4 when bubbled to saturation with 95% O2 and 5% CO2 mixture (carbogen).21 Use this ACSF for both the dissection, slice preparation and for perfusion during the electrophysiological recordings. &#160; 
	NOTE: Use clean apparatus for measuring and holding the ACSF. Using unclean apparatus may lead to cloudy solutions or formation of precipitates. Use deionized water for all the preparations.</li>
	<li>Prepare a 2 L 10x ACSF stock excluding NaHCO3 and D-Glucose in a volumetric flask. Add reagents to deionized water in the following order: NaCl (144.96g), KCl (5.52g), MgSO4.7H20 (4.92g), CaCl2.2H20 (7.56 g), KH2PO4 (3.28 g) and top up to a volume of 2 L. Stir continuously for at least 30 min&#160;using a magnetic stirrer to ensure all reagents are dissolved. Store the stock in 4 oC and use within 2 weeks.</li>
	<li>Prior to the dissection and the experiments, dilute the ACSF stock in a volumetric flask along with the addition of required amounts of NaHCO3 and D-glucose. For 1 L solution, dilute 100 ml of the stock to 1 L after adding 2.07 g NaHCO3 and 1.802 g D-glucose. The ACSF should be a clear solution free of any precipitate or undissolved particles.</li>
	<li>Cool about 200-300 ml of ACSF on ice, to be used during dissection. Ensure that the ACSF used for dissection is between 2-4 &#176;C. Use the remaining ACSF for electrophysiological experiments. Bubble all ACSF solutions to saturation with carbogen (5% CO2, 95% O2) continuously. While waiting for the ACSF to cool, prepare the dissection area and the slice chamber.</li>
</ol>
2. Preparation of Interface Chamber &#160;
NOTE: An interface brain slice chamber, used for incubating the slices and maintaining them during electrophysiological recordings (Figure 2B), consists of two compartments. The lower chamber contains distilled water maintained at 32 &#176;C by a temperature controller and continuously bubbled with carbogen.
<ol>
	<li>Switch on the temperature controller and preset it at 32 &#176;C. Wash the upper chamber for 10 to 15 min&#160;by running distilled water through the inflow tubing. Ensure that the upper chamber is clean before placing the net. Check that water level in the bottom chamber is about 70% filled with distilled water.</li>
	<li>Place the net in the upper chamber to provide a resting surface for slices (Figure 2C). Adjust the outflow tubing to ensure that the solution level is sufficiently wetting the whole area of the net. Place the lid above the net to maintain a humidified carbogen atmosphere within the upper chamber.</li>
	<li>Adjust the flow rate to 1 ml/min. Maintain this flow rate throughout the slice incubation period and the experiment. Start carbogenating the freshly prepared 1x ACSF and immerse the inflow tubing into the ACSF. Allow 20 min&#160;for the ACSF to be saturated with carbogen and for the upper chamber to be filled with it.</li>
</ol>
3. Preparation of Acute Hippocampal Slices &#160;
NOTE: The dissection protocol consists of (1) Removal of brain from the animal into cold ACSF and (2) Isolation and slicing of the hippocampus. In order for neurons to remain viable, isolate and place the brain in cold ACSF quickly and complete the whole process including slicing within 3-5 min.
<ol>
	<li>Removal of the brain into cold ACSF
	<ol>
		<li>Lay out the dissection tools in the manner shown in Figure 1A. Arrange the tools according to the order of use to facilitate the dissection process. Before starting, ensure all the dissection tools are ready.</li>
		<li>Mount a razor blade, cleaned with ethyl acetate, absolute ethanol and distilled water, onto the manual tissue chopper (Figure 1B), secure it firmly and ensure that the cutting edge is evenly aligned. Test chopping a piece of filter paper to ensure that the blade is firmly secured. Set the sliding Vernier micrometer to its starting position.</li>
		<li>Euthanize the animal using carbon dioxide (CO2) in an induction chamber and decapitate with bandage scissors or Guillotine. Using an Iris scissors, remove the skin and fur above the skull. Make a cut through the posterior to remove the brainstem.Make a small incision along the right side of the skull and a longer incision on the left. 
		CAUTION! Make only a small incision to the side that is used for the experiments to avoid damaging it. When inserting the scissors, make sure that the force applied is upwards to avoid damage to the brain.</li>
		<li>Carefully remove the skull with a bone rongeur starting from the left to the right side of skull to reveal the cortex. A thin layer of dura can also be seen. Carefully remove the frontal plates with the rongeur. Remove most of the dura along with the frontal plates. 
		CAUTION! Be careful that the dura does not slice through the brain tissue.</li>
		<li>Remove the remaining dura, if any, specifically in the junction between cortex and cerebellum with the flat end of a spatula. For steps 3.1.5 and 3.1.6, maintain pressure upwards i.e., away from the brain to avoid damaging it. Using the spatula, gently scoop the brain into a petri dish filled with cold and carbogenated ACSF (2-4 &#176;C), placed on an aluminum cooling block.</li>
	</ol>
	</li>
	<li>Isolation of the hippocampus
	<ol>
		<li>Using a scalpel, make a straight cut to remove the cerebellum and another cut to remove the anterior portion of the brain (approximately one quarter). Make a shallow cut along the midline.</li>
		<li>Carefully remove the cortex with a sickle scaler, starting from the midline to reveal the dorsal hippocampus. Remove the layer of cortex above the hippocampus. Use fingers or angled forceps to support the brain. Make a small cut to the hippocampal commissure. Gently remove the hippocampus with the sickle scaler starting from the dorsal hippocampus using rolling motions. 
		CAUTION! Be gentle to avoid stretching and tearing the hippocampus.</li>
		<li>Remove any cortex and connective tissues around the isolated hippocampus with the sickle scaler.</li>
	</ol>
	</li>
	<li>Slicing the hippocampal tissue and transferring the slices onto the interface chamber
	<ol>
		<li>Place a piece of ACSF-soaked filter paper (Grade 1, 30 mm) on the slicing stage of the manual slicer. Scoop and place the hippocampal tissue onto the filter paper. Move the filter paper to align the hippocampus at a proper orientation in relation to the blade of the slicer so that the hippocampus is sliced at an angle of about 70o to the fimbria.</li>
		<li>Blot the excess solution surrounding the hippocampal tissue with a folded filter paper (Grade 1, 85 mm) leaving the hippocampus slightly wet. Start slicing the hippocampus transversely. Slice and discard tissue from the extreme end of the hippocampus where the slice morphology is not clear.</li>
		<li>Slice the remaining tissue into 400 &#181;m-thick slices. Pick up hippocampal slices gently from the blade with a brush with soft bristles using gentle swiping movements and place the slices into a small beaker filled with cold carbogenated ACSF. Perform the steps 3.3.1-3.3.3 as quickly as possible since the hippocampal tissue is exposed to air. &#160; 
		NOTE: Generally two-thirds of the hippocampus is sliced, and 4-6 slices with clear morphology can be prepared.</li>
		<li>Transfer the slices gently onto the net in the slice chamber using a clean plastic Pasteur pipette with a broad tip (made by cutting away 2-3 cm of the tip). Carefully adjust the position of the slices on the net using a small syringe with a bent tip. Position the slices in a manner that facilitates electrode location and recording. Check to ensure that the slices are sufficiently surrounded by ACSF but are not submerged or floating (Figure 2C-D). Cover the chamber and incubate the slices for 2-3 hr. &#160; 
		NOTE: The pyramidal cell layer in the healthy slices should show some transparency.</li>
	</ol>
	</li>
</ol>
4. Recording of CA3-CA1 Synaptic Responses &#160;
NOTE: The electrophysiology set-up used for field potential recording is shown in Figure 2A. A Faraday cage is strongly recommended if the electrical interference is beyond the control after the proper grounding of the electrical settings. Many different types of submerged and interface chambers are commercially available. However, interface chambers are preferred as slices exhibit more robust synaptic responses in them.
<ol>
	<li>Positioning of electrodes
	<ol>
		<li>Turn on the electrical apparatus (stimulators and amplifiers) to be used. Mount and secure the stimulating and recording electrodes in the plexiglass holders of the micromanipulators. &#160; 
		NOTE: We use monopolar, lacquer-coated, stainless steel electrodes of 5 M&#937; resistance for both stimulating and recording purposes.</li>
		<li>Before use, insert these electrodes inside the pulled glass capillaries and secure with epoxy glue exposing only small portion of the electrode tip (Figure 2E). This gives strength to the otherwise slender electrodes and helps to secure them firmly in the electrode holders.</li>
		<li>Guided under the microscope, position the stimulating electrode(s) in the stratum radiatum of the CA1 region to stimulate the Schaffer collateral fibers and the recording electrode in the apical dendritic region of CA1 to record field-EPSP (fEPSP) responses. &#160; 
		NOTE: Approaching the liquid surface above the slice with the electrodes gives a sound that helps to locate quickly the surface of the slice (provided, the amplifier is connected to a loudspeaker).</li>
		<li>In synaptic tagging and capture experiments, according to the need of the experiment, position two or three stimulating electrodes (S1, S2 or S3) on either side of the recording electrode to stimulate two or more independent but overlapping inputs. Position the stimulating and recording electrodes about 200 &#181;m apart.</li>
		<li>If necessary, locate another recording electrode in the stratum pyramidale layer for recording population spike (Figure 3A).When both the electrodes have touched the slice, using the acquisition software, give a test stimulation to ensure a proper fEPSP signal. &#160; 
		NOTE: We use biphasic, constant-current pulses (impulse duration 0.1 msec/half-wave) for test stimulation.</li>
		<li>Once a proper fEPSP signal is obtained, carefully lower the electrodes about 200 &#181;m deep using fine movement knobs of the manipulators. Allow 20 min&#160;for the slices to recover. Test the pathway independence with a paired-pulse facilitation protocol27,28.</li>
	</ol>
	</li>
	<li>Input-output relation
	<ol>
		<li>Determine the input-output relation (afferent stimulation vs fEPSP slope) for each input by measuring the slope value at a range of current intensities. Perform this between 20 &#181;A to 100 &#181;A. Then set the stimulation intensity for each input to obtain 40% of the maximum fEPSP slope. Keep this constant throughout the experiment.</li>
		<li>After 15-20 min, start recording the baseline. Monitor the fEPSP slope closely during this period and reset the stimulus intensity if the slope fluctuates more than 10% from the set value and start a new baseline. Record at least 30 min&#160;or 1 hr stable baseline before proceeding. &#160; 
		NOTE: For the test or the baseline stimulation, we use four sweeps of 0.2 Hz biphasic, constant current pulses (0.1 msec per polarity) given every 5 min. An average slope of these four responses is then considered as one repeat. The signals are filtered and amplified by a differential amplifier, digitized using an analog-to-digital converter and monitored online with custom-made software.</li>
	</ol>
	</li>
	<li>Induction of LTP/LTD using stimulation protocols 
	NOTE: Both LTP and LTD have been classified as early and late-LTP/LTD based on the requirements of protein synthesis; the latter requiring translation and/or transcription for its late maintenance [for review see4]. A variety of electrical stimulation paradigms can specifically induce the different forms of LTP and LTD.
	<ol>
		<li>WTET: 100 Hz, 21 biphasic constant current pulses (0.2 msec per phase).</li>
		<li>STET: Three bursts of 100 pulses for one second (100 Hz) every 10 min ( Pulse width &#160;0.2 msec per phase).</li>
		<li>900 bursts over a 15 min duration . 1 burst consists of 3 pulses (0.2 msec width) with an interpulse interval of 50 msec (20 Hz).&#160; The inter burst interval is 1 sec ( total number of pulses 2,700).</li>
	</ol>
	</li>
</ol>
5. Cleaning of Slice Chamber and Perfusion System
<ol>
	<li>After the recording is over, collect the hippocampal slices for further biochemical analysis or else discard appropriately. Turn off the carbogen supply and temperature controller. Wash the carbogen bubbler in distilled water.</li>
	<li>Clean the net thoroughly with a brush and distilled water. Wash the rig for 15-20 min&#160;with distilled water at a higher flow rate. Once in 3-4 days, change the distilled water in the lower compartment of the chamber and also clean the chamber regularly with 3% hydrogen peroxide solution to avoid fungal growth.</li>
</ol>

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U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anisomycin+inhibits+the+late+maintenance+of+long-term+depression+in+rat+hippocampal+slices+in+vitro.">Anisomycin inhibits the late maintenance of long-term depression in rat hippocampal slices in vitro.</a> Neuroscience Letters. 338, 147-150 (2003).</li><li>Ishikawa, Y., Tamura, H., Shiosaka, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diversity+of+neuropsin+(KLK8)-dependent+synaptic+associativity+in+the+hippocampal+pyramidal+neuron.">Diversity of neuropsin (KLK8)-dependent synaptic associativity in the hippocampal pyramidal neuron.</a> The Journal of physiology. 589, 3559-3573 (2011).</li><li>Ishikawa, Y., Horii, Y., Tamura, H., Shiosaka, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuropsin+(KLK8)-dependent+and+-independent+synaptic+tagging+in+the+Schaffer-collateral+pathway+of+mouse+hippocampus.">Neuropsin (KLK8)-dependent and -independent synaptic tagging in the Schaffer-collateral pathway of mouse hippocampus.</a> The Journal of neuroscience : the official journal of the Society for Neuroscience. 28, 843-849 (2008).</li><li>Sajikumar, S., Morris, R. G. M., Korte, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Competition+between+recently+potentiated+synaptic+inputs+reveals+a+winner-take-all+phase+of+synaptic+tagging+and+capture.">Competition between recently potentiated synaptic inputs reveals a winner-take-all phase of synaptic tagging and capture.</a> Proceedings of the National Academy of Sciences. 111, 12217-12221 (2014).</li><li>Navakkode, S., Sajikumar, S., Frey, J. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+type+IV-specific+phosphodiesterase+inhibitor+rolipram+and+its+effect+on+hippocampal+long-term+potentiation+and+synaptic+tagging.">The type IV-specific phosphodiesterase inhibitor rolipram and its effect on hippocampal long-term potentiation and synaptic tagging.</a> The Journal of neuroscience : the official journal of the Society for Neuroscience. 24, 7740-7744 (2004).</li><li>Kohl, M. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hemisphere-specific+optogenetic+stimulation+reveals+left-right+asymmetry+of+hippocampal+plasticity.">Hemisphere-specific optogenetic stimulation reveals left-right asymmetry of hippocampal plasticity.</a> Nature. 14, 1413-1415 (2011).</li><li>Frey, U., Huang, Y. Y., Kandel, E. R. <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+cAMP+Simulate+a+Late+Stage+of+LTP+in+Hippocampal+CA1+Neurons.">Effects of cAMP Simulate a Late Stage of LTP in Hippocampal CA1 Neurons.</a> Science. 260, 1661-1664 (1993).</li><li>Erdogdu, G., Uto, A., Hossmann, K. -. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+of+global+ischemia+and+recirculation+of+rat+brain+on+protein+synthesis+in+vitro.">The effect of global ischemia and recirculation of rat brain on protein synthesis in vitro.</a> Metab Brain Dis. 8, 199-206 (1993).</li><li>Djuricic, B., Berger, R., Paschen, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+synthesis+and+energy+metabolism+in+hippocampal+slices+during+extended+(24+hours)+recovery+following+different+periods+of+ischemia.">Protein synthesis and energy metabolism in hippocampal slices during extended (24 hours) recovery following different periods of ischemia.</a> Metab Brain Dis. 9, 377-389 (1994).</li><li>Park, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA+receptor-dependent+long-term+potentiation+comprises+a+family+of+temporally+overlapping+forms+of+synaptic+plasticity+that+are+induced+by+different+patterns+of+stimulation.">NMDA receptor-dependent long-term potentiation comprises a family of temporally overlapping forms of synaptic plasticity that are induced by different patterns of stimulation.</a> Philosophical Transactions of the Royal Society B: Biological Sciences. 369, 20130131 (2014).</li></ol>

Open access for this video article is sponsored by Cerebos Pacific Limited.

Acute hippocampal slice is an excellent model system for the study of LTP and other functional plasticity processes such as STC and cross-capture. It preserves much of the laminar structural network of the hippocampal circuits, allows precise electrode locations and offers alongside, an open platform for rapid neuropharmacological manipulation without a blood-brain barrier.
This article describes the methodology for the preparation of viable acute hippocampal slices from young adult rats and using them to investigate STC and cross-tagging. Previous research has emphasized that gender and age of the animals are important factors to consider for use in electrophysiology studies.27,28 Therefore young adult animals with fully expressed adult receptor functions (male Wistar rats aged 5-7 weeks) are used.23 Asymmetries in the connections between the left and right hippocampus have been noted in rodents29and major differences in NMDA receptor expression have been reported as well 34. We have used the right hippocampus in order to be consistent with our previous LTP studies.23,32 However, either of the hippocampi can be used as long as consistency is maintained.
As in any protocol, it is very crucial to perform the isolation and slicing procedures quickly but taking care that the tissue is not stretched, damaged, rendered dry or hypoxic. The variations in pH, temperature and ionic composition of the solutions can have profound effect on the viability of the slices and the results. Hence such variations should be avoided. It has been observed that glutamate receptor-dependent calcium release occurring during the preparation steps can irreversibly affect protein synthesis in nervous tissue 35,36,37. Using manual tissue slicers can help to minimize this by allowing the process to be completed very quickly as compared to vibraslicers. However, many laboratories also effectively use vibraslicers with necessary precautions to preserve slice viability. Another important factor to consider is the long incubation period before starting the experiments. This has been noted to be really crucial to achieve stability in metabolic state and kinase activation levels in the slices after the disturbance caused during preparation 23. Such stability is necessary for consistency in long-term recordings. We re-emphasize on this observation and suggest the long incubation hours of about 3 hr.
A variety of stimulation parameters are known to induce LTP, but the molecular mechanisms elicited in each case may not be the same (for review see 38). This can influence the durability and other characteristics of the LTP which, in turn, can affect the results of synaptic tagging and capture experiments. Hence it is important to validate the stimulation paradigms and characteristics of the elicited LTP under the conditions of the performing laboratory and maintain consistency.
We generally do not consider experiments with very large presynaptic fiber volleys and with maximal fEPSPs less than 0.5 mV and the experiments involving substantial changes in the fiber volley during the recordings are also rejected. Further, while performing two-pathway or three-pathway experiments, it is important to ensure the pathway independence. This can be carried out with a paired-pulse facilitation protocol28.
One downside of the interface recording systems is the formation of condensation droplets on the electrodes during the long recording hours due to the temperature and humidity differences between the chamber and the surroundings. These droplets need to be carefully blotted from time to time. Otherwise the droplets can drip onto the slices and cause disturbance or even loss of signals. We usually tackle this by skilfully blotting the droplets guided under the microscope using a slender filter paper wick, without touching the electrodes. However, the best solution would be to use a centralized heating system, such as the ETC system developed by University of Edinburgh researchers.
On a concluding note, a variety of methodologies exist in the laboratories worldwide that are used for the preparation of hippocampal slices for different experimental purposes. Each of the procedure offers some advantages over the other. One needs to carefully optimize the minute details of the protocol to suit the purpose of the experiment. We hope that this article helps in improving some aspects of the methodology for studying late-associative processes such as STC and cross-capture.

The encoding and storage of information in the brain still remains the most significant and keenly pursued challenge in neuroscience. Over the years, long-term potentiation (LTP) and long-term depression (LTD) have emerged as the leading cellular correlates of memory1,2. These activity dependent changes, which exhibit input specificity and associativity, result in the stabilization of memory traces in the neuronal networks 1,3,4. The maintenance of the two forms of synaptic plasticity requires the synthesis of plasticity-related products (PRPs)5-10. Synapse specificity that involves the interaction of newly synthesized protein only with specific activated synapses expressing LTP or LTD, is critical to memory. This specificity is explained by the concept of &#8216;Synaptic Tagging and Capture&#8217; (STC), where the PRPs interact with recently active, &#8216;tagged&#8217; synapses11,12. The STC process offers a framework for associative properties of memories at the cellular level. It provides us with a conceptual basis of how short-term forms of plasticity are transformed into long-lasting forms of plasticity in an associative and time-dependent manner13.
During the process of STC, a strong tetanization in one input that leads to protein synthesis dependent late-LTP, results in the reinforcement of a protein synthesis independent early-LTP induced in another independent input on to the same population of neurons into a persistent one13. The setting of a local synaptic tag by a transient neural activity and the synthesis of the diffusible PRPs by the strong neural activity are the two key events during STC13,14. The capture of the PRPs by the recently potentiated &#8216;tagged&#8217; synapses is fundamental to the maintenance of long-term potentiation. Many studies have been done to confirm the existence of STC phenomenon15-17 and identify the candidate &#8216;tags&#8217;18 and &#8216;PRPs&#8217;19. Calcium/calmodulin-dependent protein kinase II (CaMKII) and extracellular signal-regulated kinase1/2 (ERK1/2); CaMKIV, Protein Kinase M (PKM) and brain-derived neurotrophic factor (BDNF) are some of the candidate molecules for &#8216;tag&#8217; and &#8216;PRP&#8217; respectively19-21. The synaptic tagging model has further been expanded to include the positive associative interactions between LTP and LTD - the &#8220;synaptic cross-tagging&#8221;22. In synaptic cross-tagging, a late LTP/ LTD in one synaptic input transforms the opposite protein synthesis-independent early-LTD/LTP in an independent input into its long-lasting form or vice versa22. 
The hippocampal slice preparation is the most widely used model in the studies of long-term synaptic plasticity23,24. Much of the understanding about the concepts of synaptic tagging and cross- tagging arises from the studies done in CA1 region of the hippocampus and because of the technical complexity many of the labs are still unable to study these processes. Experimental conditions for the preparation of rat hippocampal slices and the recording of stable late-LTP/LTD for extended hours are extremely important to study synaptic tagging/cross-tagging23,25,26. This article describes the detailed experimental procedures for studying long-term plasticity processes such as STC and cross-tagging in the CA1 pyramidal neurons using stable, long-term field-potential recordings from acute hippocampal slices of rats.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>I. Dissection Tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1. Bandage scissors</td>
			<td>KLS Martin, Germany</td>
			<td>21-195-23-07</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>B-Braun/Aesculap, Germany</td>
			<td>LX553R</td>
			<td></td>
		</tr>
		<tr>
			<td>2. Iris scissors</td>
			<td>B-Braun/Aesculap, Germany</td>
			<td>BC140R,</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>BC100R</td>
			<td></td>
		</tr>
		<tr>
			<td>3. Bone rongeur</td>
			<td>World Precision Instruments (WPI), Germany</td>
			<td>14089-G</td>
			<td></td>
		</tr>
		<tr>
			<td>4. Scalpel</td>
			<td>World Precision Instruments (WPI), Germany;</td>
			<td>500236-G</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td>B-Braun/Aesculap, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>BB73</td>
			<td></td>
		</tr>
		<tr>
			<td>5. Scalpel blade#11</td>
			<td>B-Braun/Aesculap, Germany</td>
			<td>BB511</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>6. Sickle scaler</td>
			<td>KLS Martin, Germany</td>
			<td>38-685-00</td>
			<td></td>
		</tr>
		<tr>
			<td>7. Angled forceps</td>
			<td>B-Braun/Aesculap, Germany</td>
			<td>BD321R</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>8. Anesthetizing/Induction chamber</td>
			<td>MIP Anesthesia Technologies (Now, Patterson Scientific), Oregon</td>
			<td>AS-01-0530-LG</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
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			<td>II. ACSF component chemicals</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1. Sodium chloride (NaCl)</td>
			<td>Sigma-Aldrich</td>
			<td>S5886</td>
			<td></td>
		</tr>
		<tr>
			<td>2. Potassium chloride (KCl)</td>
			<td>Sigma-Aldrich</td>
			<td>P9541</td>
			<td></td>
		</tr>
		<tr>
			<td>3. Magnesium sulphate heptahydrate (MgSO4.7H20)</td>
			<td>Sigma-Aldrich</td>
			<td>M1880</td>
			<td></td>
		</tr>
		<tr>
			<td>4. Calcium chloride dihydrate (CaCl2.2H2O)</td>
			<td>Sigma-Aldrich</td>
			<td>C3881</td>
			<td></td>
		</tr>
		<tr>
			<td>5. Potassium phosphate monobasic (KH2PO4)</td>
			<td>Sigma-Aldrich</td>
			<td>P9791</td>
			<td></td>
		</tr>
		<tr>
			<td>6. Sodium bicarbonate (NaHCO3)</td>
			<td>Sigma-Aldrich</td>
			<td>S5761</td>
			<td></td>
		</tr>
		<tr>
			<td>7. D-Glucose anhydrous (C6H12O6)</td>
			<td>Sigma-Aldrich</td>
			<td>G7021</td>
			<td></td>
		</tr>
		<tr>
			<td>III. Electrophysiology Instruments</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1. Microscope</td>
			<td>Olympus, Japan</td>
			<td>Model SZ61</td>
			<td></td>
		</tr>
		<tr>
			<td>2. Temperature Controller</td>
			<td>Scientific Systems Design Inc. Canada</td>
			<td>PTC03</td>
			<td></td>
		</tr>
		<tr>
			<td>3. Differential AC Amplifier</td>
			<td>AM Systems, USA</td>
			<td>Model 1700</td>
			<td></td>
		</tr>
		<tr>
			<td>4. Isolated Pulse Stimulator</td>
			<td>AM Systems, USA</td>
			<td>Model 2100</td>
			<td></td>
		</tr>
		<tr>
			<td>5. Oscilloscope</td>
			<td>Rhode &#38; Schwarz</td>
			<td>HM0722</td>
			<td></td>
		</tr>
		<tr>
			<td>6. Digital-Analog Converter</td>
			<td>Cambridge Electronic Design Ltd. Cambridge, UK</td>
			<td>CED-Power 1401-3</td>
			<td></td>
		</tr>
		<tr>
			<td>7. Interface Brain Slice Chamber</td>
			<td>Scientific Systems Design Inc. Canada</td>
			<td>BSC01</td>
			<td></td>
		</tr>
		<tr>
			<td>8. Tubing Pump</td>
			<td>Ismatec, Idex Health &#38; Science, Germany</td>
			<td>REGLO-Analog</td>
			<td></td>
		</tr>
		<tr>
			<td>9. Carbogen Flowmeter</td>
			<td>Cole-Parmer</td>
			<td>03220-44</td>
			<td></td>
		</tr>
		<tr>
			<td>10. Fiber Light Illuminator</td>
			<td>Dolan-Jenner Industries</td>
			<td>Fiber Lite MI-150</td>
			<td></td>
		</tr>
		<tr>
			<td>11. Micromanipulators</td>
			<td>Marzhauser Wetzlar, Germany</td>
			<td>00-42-101-0000 (MM-33)</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>00-42-102-0000 (MM-32)</td>
			<td></td>
		</tr>
		<tr>
			<td>12. Electrodes</td>
			<td>AM Systems, USA</td>
			<td>571000 (Stainless steel; 0.010, 5M&#8486;, 8 degree)</td>
			<td></td>
		</tr>
	</tbody>
</table>

investigation of synaptic tagging/capture and cross-capture using acute hippocampal slices from rodents

Synaptic tagging and capture (STC) and cross-tagging are two important mechanisms at cellular level that explain how synapse-specificity and associativity is achieved in neurons within a specific time frame. These long-term plasticity-related processes are the leading candidate models to study the basis of memory formation and persistence at the cellular level. Both STC and cross-tagging involve two serial processes: (1) setting of the synaptic tag as triggered by a specific pattern of stimulation, and (2) synaptic capture, whereby the synaptic tag interacts with newly synthesized plasticity-related proteins (PRPs). Much of the understanding about the concepts of STC and cross-tagging arises from the studies done in CA1 region of the hippocampus and because of the technical complexity many of the laboratories are still unable to study these processes. Experimental conditions for the preparation of hippocampal slices and the recording of stable late-LTP/LTD are extremely important to study synaptic tagging/cross-tagging. This video article describes the experimental procedures to study long-term plasticity processes such as STC and cross-tagging in the CA1 pyramidal neurons using stable, long-term field-potential recordings from acute hippocampal slices of rats.

The described methodology has been used to&#160;study long-lasting forms of LTP/LTD and&#160; its associative interactions such as synaptic tagging&#160; and cross-capture&#160;&#160;from&#160;acute hippocampal slices of adult&#160; rats.23&#160;This technique has proven effective for experiments with both rats (Wistar) and a variety of mouse strains30,31. The methodology has been used successfully for stable LTP recordings of up to 8-12 hr.32
The &#8216;tag&#8217; set by the weak tetanization of one input (S1) captures the &#8216;PRPs&#8217; induced by the strong tetanization of another independent but overlapping input (S2; Figure 3B, filled circles) thereby transforming the otherwise decaying form of LTP (early-LTP) in S1 into a long-lasting one (Figure 3B, open circles) (For comparison of early-LTP induced by WTET see20,33). The PRPs captured by the weak tetanization set tag need not necessarily come from the STET-induced late-LTP but can also be provided by the SLFS-induced late-LTD. This type of positive associative interaction between LTP and LTD is referred to as &#8216;cross-tagging/capture&#8217;. The WTET-induced early-LTP in S1 gets reinforced to late-LTP (Figure 3C, open circles) by capturing the PRPs provided by the SLFS-induced late-LTD in S2 (Figure 3C, filled circles). Statistically significant potentiation or depression was &#160;&#160;maintained in S1 and S2 in both cases when compared to its&#160;own baseline (Wilcoxon test;&#160;P&#160;&#60; 0.05).&#160;
For the tag-PRP interaction to occur, the temporal order of the two events (weak-before-strong/strong-before-weak) is not crucial as long as the time window between the two events remains within the range of 30-60 minutes. It would be wise to include a third, independent but overlapping synaptic input and use it as a baseline control to monitor the stability of recordings. The electrical stimulation protocols used to induce early- and late forms of LTP/LTD must be validated in single-input experiments for consistency and reliability before using them in STC experiments. We would also like to emphasize the importance of slice preparation methodology described in the protocol since the success of these experiments relies heavily on the quality of the slices.
<img alt="Figure 1" src="/files/ftp_upload/53008/53008fig1.jpg" /> 
Figure 1.&#160;(A) Tools used in the dissection of hippocampus: (a) Bandage Scissors (b) Iris scissors (c) Bone rongeur (d) Thin spatula, (e) Scalpel number 11 (f) Sickle scaler (g) Soft-bristle paint brush (h) Plastic Pasteur pipette (i) filter paper (85mm) (j) filter paper (30mm) (k) Glass beakers (l) Aluminum cooling blocks to fit petri dish and beakers (m) Petri dish. (B) Manual tissue chopper. (a) Platform (b) Cutting arm with blade-holder (c) Vernier micrometer, resolution 10 microns. <a href="https://www.jove.com/files/ftp_upload/53008/53008fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53008/53008fig2.jpg" /> 
Figure 2.&#160;Electrophysiology set-up for field-potential recordings consisting of (A) stimulators (b) a differential amplifier (c) an analog-to-digital converter (d) Oscilloscope (e) computer with acquisition software (f) Vibration-resistant table-top (g) microscope with &#62;4x magnification (h) interface brain-slice chamber (i) a perfusion system for ACSF and carbogen supply (j) temperature controller (k) an illumination source (l) manipulators with electrode holders. (B) Interface brain-slice chamber. (C) &#38; (D) Hippocampal slices in the interface chamber. (E) Stainless steel electrode sealed in a glass capillary. <a href="https://www.jove.com/files/ftp_upload/53008/53008fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53008/53008fig3.jpg" /> 
Figure 3.&#160;(A) Schematic representation of a transverse hippocampal slice and electrode location for field-potential recording: In this representation, two stimulating electrodes (S1 and S2) are positioned in the stratum radiatum of the CA1 region to stimulate two independent but overlapping synaptic inputs onto CA1 pyramidal neurons. Two extracellular recording electrodes, one to record field-EPSP (excitatory post-synaptic potential) from the apical dendritic compartment and another to record somatic population spike from the pyramidal cell bodies, are located in the stratum radiatum and stratum pyramidale respectively. CA1- cornu ammonis region 1, CA3- cornu ammonis region 3, DG- dentate gyrus, SC- Schaffer collateral fibers, S1- stimulating electrode 1, S2-stimulating electrode 2. (B) Weak before strong paradigm to study STC: Weak tetanization (WTET) is applied to S1 (open circles) for inducing early-LTP followed by strong tetanization (STET) of S2 (filled circles) at 30 min&#160;to induce late-LTP. The early-LTP in S1 gets reinforced to late-LTP showing tagging and capture interaction (n = 6). (C) Weak before strong paradigm to study cross-tagging: Early-LTP is induced by WTET in S1 (open circles) followed by the induction of late-LTD in S2 (filled circles) using SLFS after 30 min. In S1, the early-LTP is transformed to late-LTP lasting 6 hr showing cross-tagging and capture (n = 6). Single arrow represents weak tetanization applied for inducing early-LTP. Triplet of arrows represents strong tetanization for inducing late-LTP. The broken arrow represents the time point at which SLFS was applied to the representative synaptic input to induce late-LTD. Error bars indicate SEM. <a href="https://www.jove.com/files/ftp_upload/53008/53008fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Investigation of Synaptic Tagging/Capture and Cross-capture using Acute Hippocampal Slices from Rodents at JoVE.com

Investigation of Synaptic Tagging/Capture and Cross-capture using Acute Hippocampal Slices from Rodents

This video article describes experimental procedures to study long-term plasticity and its associative processes such as synaptic tagging, capture and cross-tagging in the CA1 pyramidal neurons using acute hippocampal slices from rodents.

Synaptic tagging and capture (STC) and cross-tagging are two important mechanisms at cellular level that explain how synapse-specificity and associativity ...

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13.7K Views.  National University of Singapore. This video article describes experimental procedures to study long-term plasticity and its associative processes such as synaptic tagging, capture and cross-tagging in the CA1 pyramidal neurons using acute hippocampal slices from rodents.

Video: Investigation of Synaptic Tagging/Capture and Cross-capture using Acute Hippocampal Slices from Rodents

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 ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

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 ...

Post-embedding Immunogold Labeling of Synaptic Proteins in Hippocampal Slice Cultures

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as colloidal gold can reveal the localization and distribution of specific antigens in various tissues1. The two most widely used techniques are pre-embedding and post-embedding techniques. In pre-embedding immunogold-electron microscopy (EM) techniques, the tissue must be permeabilized to allow antibody penetration before it is embedded. These techniques are ideal for preserving structures but poor penetration of the antibody (often only the first few micrometers) is a considerable drawback2. The post-embedding labeling methods can avoid this problem because labeling takes place on sections of fixed tissues where antigens are more easily accessible. Over the years, a number of modifications have improved the post-embedding methods to enhance immunoreactivity and to preserve ultrastructure3-5.
Tissue fixation is a crucial part of EM studies. Fixatives chemically crosslink the macromolecules to lock the tissue structures in place. The choice of fixative affects not only structural preservation but also antigenicity and contrast. Osmium tetroxide (OsO4), formaldehyde, and glutaraldehyde have been the standard fixatives for decades, including for central nervous system (CNS) tissues that are especially prone to structural damage during chemical and physical processing. Unfortunately, OsO4 is highly reactive and has been shown to mask antigens6, resulting in poor and insufficient labeling. Alternative approaches to avoid chemical fixation include freezing the tissues. But these techniques are difficult to perform and require expensive instrumentation. To address some of these problems and to improve CNS tissue labeling, Phend et al. replaced OsO4 with uranyl acetate (UA) and tannic acid (TA), and successfully introduced additional modifications to improve the sensitivity of antigen detection and structural preservation in brain and spinal cord tissues7. We have adopted this osmium-free post-embedding method to rat brain tissue and optimized the immunogold labeling technique to detect and study synaptic proteins.
We present here a method to determine the ultrastructural localization of synaptic proteins in rat hippocampal CA1 pyramidal neurons. We use organotypic hippocampal cultured slices. These slices maintain the trisynaptic circuitry of the hippocampus, and thus are especially useful for studying synaptic plasticity, a mechanism widely thought to underlie learning and memory. Organotypic hippocampal slices from postnatal day 5 and 6 mouse/rat pups can be prepared as described previously8, and are especially useful to acutely knockdown or overexpress exogenous proteins. We have previously used this protocol to characterize neurogranin (Ng), a neuron-specific protein with a critical role in regulating synaptic function8,9 . We have also used it to characterize the ultrastructural localization of calmodulin (CaM) and Ca2+/CaM-dependent protein kinase II (CaMKII)10. As illustrated in the results, this protocol allows good ultrastructural preservation of dendritic spines and efficient labeling of Ng to help characterize its distribution in the spine8. Furthermore, the procedure described here can have wide applicability in studying many other proteins involved in neuronal functions.

The localization and distribution of proteins provide important information for understanding their cellular functions. The superior spatial resolution of electron microscopy (EM) can be used to determine the subcellular localization of a given antigen following immunohistochemistry. For tissues of the central nervous system (CNS), preserving structural integrity while maintaining antigenicity has been especially difficult in EM studies. Here, we adopt a procedure that has been used to preserve structures and antigens in the CNS to study and characterize synaptic proteins in rat hippocampal CA1 pyramidal neurons.

Immunoelectron microscopy is a powerful tool to study biological molecules at the subcellular level. Antibodies coupled to electron-dense markers such as ...

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 ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic ...

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 ...

Evaluation of Synapse Density in Hippocampal Rodent Brain Slices

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is tightly regulated during development and neuronal maturation. Importantly, deficits in synapse number can lead to cognitive dysfunction. Therefore, the evaluation of synapse number is an integral part of neurobiology. However, as synapses are small and highly compact in the intact brain, the assessment of absolute number is challenging. This protocol describes a method to easily identify and evaluate synapses in hippocampal rodent slices using immunofluorescence microscopy. It includes a three-step procedure to evaluate synapses in high-quality confocal microscopy images by analyzing the co-localization of pre- and postsynaptic proteins in hippocampal slices. It also explains how the analysis is performed and gives representative examples from both excitatory and inhibitory synapses. This protocol provides a solid foundation for the analysis of synapses and can be applied to any research investigating the structure and function of the brain.

A protocol for accurately identifying and analyzing synapses in hippocampal slices using immunofluorescence is outlined in this article.

In the brain, synapses are specialized junctions between neurons, determining the strength and spread of neuronal signaling. The number of synapses is ...

Single Synapse Indicators of Glutamate Release and Uptake in Acute Brain Slices from Normal and Huntington Mice

Synapses are highly compartmentalized functional units that operate independently on each other. In Huntington&#39;s disease (HD) and other neurodegenerative disorders, this independence might be compromised due to insufficient glutamate clearance and the resulting spill-in and spill-out effects. Altered astrocytic coverage of the presynaptic terminals and/or dendritic spines as well as a reduced size of glutamate transporter clusters at glutamate release sites have been implicated in the pathogenesis of diseases resulting in symptoms of dys-/hyperkinesia. However, the mechanisms leading to the dysfunction of glutamatergic synapses in HD are not well understood. Improving and applying synapse imaging we have obtained data shedding new light on the mechanisms impeding the initiation of movements. Here, we describe the principle elements of a relatively inexpensive approach to achieve single synapse resolution by using the new genetically encoded ultrafast glutamate sensor iGluu, wide-field optics, a scientific CMOS (sCMOS) camera, a 473 nm laser and a laser positioning system to evaluate the state of corticostriatal synapses in acute slices from age appropriate healthy or diseased mice. Glutamate transients were constructed from single or multiple pixels to obtain estimates of i) glutamate release based on the maximal elevation of the glutamate concentration [Glu] next to the active zone and ii) glutamate uptake as reflected in the time constant of decay (TauD) of the perisynaptic [Glu]. Differences in the resting bouton size and contrasting patterns of short-term plasticity served as criteria for the identification of corticostriatal terminals as belonging to the intratelencephalic (IT) or the pyramidal tract (PT) pathway. Using these methods, we discovered that in symptomatic HD mice ~40% of PT-type corticostriatal synapses exhibited insufficient glutamate clearance, suggesting that these synapses might be at risk to excitotoxic damage. The results underline the usefulness of TauD as a biomarker of dysfunctional synapses in Huntington mice with a hypokinetic phenotype.

We present a protocol to evaluate the balance between glutamate release and clearance at single corticostriatal glutamatergic synapses in acute slices from adult mice. This protocol uses the fluorescent sensor iGluu for glutamate detection, a sCMOS camera for signal acquisition and a device for focal laser illumination.