This work was supported by core facilities of the Center for Translational Neuroscience supported by NIH award P20 GM103425 and P30 GM110702 to Dr. Garcia-Rill. This work was also supported by grants from FONCYT-Agencia Nacional de Promoci&#242;n Cient&#236;fica y Tecnol&#242;gica; BID 1728 OC.AR. PICT-2012-1769 and UBACYT 2014-2017 #20120130101305BA (to Dr. Urbano).

All experimental protocols were approved by the Institutional Animal Care and Use Committee of the University of Arkansas for Medical Sciences (Protocol number #3593) and were in agreement with the National Institutes of Health guidelines for the care and use of laboratory animals.
1. Preparation of Standard-artificial Cerebrospinal Fluid (aCSF)
<ol>
	<li>Preparation of Stock Solution A
	<ol>
		<li>Add 700 ml of distilled water to a clean 1 L beaker before adding chemicals.</li>
		<li>While continuously stirring a volume of 500 ml, add 136.75 g of NaCl, 6.99 g of KCl, 2.89 g of MgSO4, and 2.83 g of NaH2PO4.</li>
		<li>Add more distilled water to reach a final volume of 1 L. After dilution, the final concentration of each compound is 117 mM NaCl, 4.69 mM KCl, 1.2 mM MgSO4, and 1.18 mM NaH2PO4.</li>
		<li>Keep refrigerated at 4 &#176;C for up to 2 weeks.</li>
	</ol>
	</li>
	<li>Preparation of Stock Solution B
	<ol>
		<li>Add 700 ml of distilled water to a clean 1L beaker before adding chemicals.</li>
		<li>While continuously stirring a volume of 500 ml, add 41.45 g of D-glucose and 41.84 g of NaHCO3.</li>
		<li>Add more distilled water to reach a final volume of 1 L. After dilution, the final concentration of each compound is 11.5 mM D-glucose and 24.9 mM NaHCO3.</li>
		<li>Keep refrigerated at 4 &#176;C for up to 2 weeks.</li>
		<li>Before each experiment mix 50 ml of stock A with 50 ml stock B and bring to 1 L with distilled water to obtain final concentration aCSF solution and leave at RT&#160;while continuously oxygenating it with carbogen (95% O2 - 5% CO2 mix) for at least 30 min. Prepare final concentration aCSF only at the beginning of each experiment and discard at the end of the day.</li>
	</ol>
	</li>
</ol>
2. Preparation of Sucrose-artificial Cerebrospinal Fluid (Sucrose-aCSF)
<ol>
	<li>Preparation of Stock Solution C
	<ol>
		<li>Add 700 ml of distilled water to a clean 1 L beaker before adding chemicals.</li>
		<li>While continuously stirring 500 ml of distilled water, add 240 g of sucrose, 6.55 g of NaHCO3, 0.671 g of KCl, 4.88 g of MgCl2, 0.22 g of CaCl2 and 0.21 g of ascorbic acid.</li>
		<li>Add more distilled water to reach a final volume of 1 L. After dilution, the final concentration of each compound is 701.1 mM sucrose, 78 mM NaHCO3, 9 mM KCl, 24 mM MgCl2, 1.5 mM CaCl2 and 1.2 mM ascorbic acid.</li>
		<li>Keep stock solution C at 4 &#176;C for up to one week.</li>
		<li>Before each experiment mix 300 ml of 50 ml of stock C with 600 ml distilled water to obtain the sucrose-aCSF solution at the final concentration and leave at RT while continuously oxygenating it with carbogen (95% O2 - 5% CO2 mix) for at least 30 min.</li>
	</ol>
	</li>
</ol>
3. Slice Preparation
<ol>
	<li>Place a clean beaker with 100 ml sucrose-aCSF solution in ice while oxygenating it with carbogen and fix the pH at 7.4 using a pH meter while adding few drops of 0.1 M NaOH solution (when pH &#60; 7.4) or 0.1 M HCl solution (when pH &#62; 7.4).</li>
	<li>Fill up a cutting chamber of a vibratome with sucrose-aCSF and oxygenate it. Turn on the glycerol-based refrigerating system coupled to the cutting chamber and wait 15 min to allow it to cool down to 0 - 4 &#176;C.</li>
	<li>Anesthetize rat pups (aged 9 to 12 from adult timed-pregnant Sprague-Dawley rats) with Ketamine (70 mg/kg, i.p.; using &#60; 50 &#181;l final volume). When pup is calm, double check that tail pinch reflex is absent.</li>
	<li>Decapitate pups.
	<ol>
		<li>Cut the head skin longitudinally from the front to back using a carbon steel scalpel blade, and pull the skin to the sides using forceps. Cut the bone covering the brain moving the scissors laterally and totally remove it to expose the brain.</li>
		<li>Then, rapidly remove the brain using a spatula (initially placed under the olfactory bulb in order to gently push out the brain from the most rostral towards the most caudal areas). Gently push the brain into ice-cooled oxygenated sucrose-artificial cerebrospinal fluid (sucrose-aCSF).</li>
	</ol>
	</li>
	<li>Make a parasagittal cut on the right hemisphere (removing approximately one third of the hemisphere), and glue the trimmed side of the brain onto a metallic disk that will be magnetically fixed to the cutting chamber of a vibroslicer to cut sagittal 400 &#181;m sections containing the pedunculopontine nucleus (PPN). Keep PPN slices at RT for 45 min prior to whole-cell patch-clamp recordings.</li>
</ol>
4. Recordings Gamma-band Oscillations in PPN Slices
<ol>
	<li>Preparation of Potassium-gluconate Intracellular Solution (High-K+ Solution)
	<ol>
		<li>Place in ice a clean beaker with 10 ml distilled water. While continuously stirring add: K+-gluconate, 90.68 mg phosphocreatine di Tris salt, 47.66 mg HEPES, 1.5 mg EGTA, 40.58 mg Mg2+-ATP, and 4 mg Na+2-GTP.</li>
		<li>Adjust pH to 7.3 with KOH (100 mM in distilled water). Add distilled water to reach a final volume of 20 mL. If needed, adjust osmolarity with sucrose (100 mM in distilled water) to be 280 - 290 mOsm. After dilution, the final concentration of each compound is 124 mM K+-gluconate, 10 mM phosphocreatine di Tris salt, 10 mM HEPES, 0.2 mM EGTA, 4 mM Mg2+-ATP, and 0.3 mM Na+2-GTP.</li>
		<li>Aliquot intracellular solutions in 1 ml tubes and freeze at -20 &#176;C. Use one aliquot per day and keep at 4 &#176;C during experiments.</li>
	</ol>
	</li>
	<li>Whole-cell Patch Clamp Recordings
	<ol>
		<li>Place slices in an immersion chamber and perfuse them (1.5 ml/min) with oxygenated (95% O2 -&#160; 5% CO2) aCSF containing the following receptor antagonists: selective NMDA receptor antagonist 2-amino-5-phosphonovaleric acid (APV, 40 &#956;M), competitive AMPA/Kainate glutamate receptor antagonist 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX, 10&#956;M), glycine receptor antagonist strychnine (STR, 10 &#956;M), the specific GABAA receptor antagonist gabazine (GBZ, 10 &#956;M), and sodium channel blocker tetrodotoxin (TTX, 3&#956;M) 
		NOTE: In the results and figures, these antagonists are collectively referred to as synaptic blockers or SBs.</li>
		<li>Fill the recording patch pipettes (Resistance 2-7 M&#937;; made from regular, commercially available thick wall borosilicate glass capillaries of 1.0 mm outer diameter and 0.6 mm inner diameter) with intracellular high-K+ solution using commercially available patch-pipette fillers with a solution filter. Insert the pipette in its amplifier&#39;s holder. Apply a small positive pressure using a 1 ml syringe connected to the pipette holder using a silicon tube connected to a three-way valve. Connect the back of the pipette holder to a patch clamp amplifier.</li>
		<li>Move the recording pipette using a mechanical micromanipulator near the PPN nucleus using a 4X objective combined to near-infrared differential interference contrast optics. 
		NOTE: PPN nucleus can be observed in slices dorsal to the superior cerebellar peduncle (SCP; observable as a thick bundle of axons). Recording pipettes were located in the PPN pars compacta, which is located immediately dorsal to the posterior end of the peduncle.</li>
		<li>Bring the recording pipette in contact with a PPN neuron while visualized using a 40X water immersion lens, and rapidly apply negative suction to form a seal with the cell.</li>
		<li>Use voltage-clamp seal software to monitor pipette resistance during negative suction using manufacturer&#39;s protocol.
		<ol>
			<li>When negative suction is slowly increased and resistance values reading by the patch-clamp monitor at the tip of the pipette reach 80 - 100 MOhm, rapidly change the holding potential to -50 mV and release the negative pressure. Start continuously applying negative suction until rupturing the neuron&#39;s membrane and electrical access is achieved in the whole-cell configuration.</li>
			<li>If access resistance values measured by the voltage-clamp seal software are 10 MOhm or higher, then continue applying negative suction in smaller amounts.</li>
		</ol>
		</li>
		<li>Compensate capacitance (i.e., slow and fast transients observed after rupturing the membrane of the cell) and series resistance in voltage-clamp mode. Switch recording mode to current clamp, and rapidly compensate bridge values (e.g., move amplifier knob or click on the automatic compensation menu using computer&#180;s mouse).
		<ol>
			<li>Continuously monitor resting membrane potential of PPN neuron being recorded using manufacturer&#39;s protocol. If resting membrane potential shift towards depolarizing or hyperpolarizing values, then use small amounts of direct current (up to 100 pA) to keep the optimum -50 mV final value. 
			NOTE: Keep only PPN neurons with a stable resting membrane potential (RMP) of -48 mV or more hyperpolarized (i.e., RMP &#60; -48 mV).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

PPN neurons have intrinsic properties that allow them to fire action potentials at beta/gamma band frequencies during in vivo recordings from animals that are awake or during REM sleep, but not during slow wave sleep2,3,5,13-17. Other authors have showed that brainstem transections at more anterior levels than PPN reduced gamma frequencies during EEG recordings. However, when brainstem lesions posterior to where this nucleus is located, the direct stimulation of PPN allowed the manifestation of cortic...

PPN nucleus is anatomically included in the caudal mesencephalic tegmentum. The PPN is a key component of RAS1. The PPN participates in the maintenance of behavioral activated states (i.e., waking, REM sleep)2. Electrical stimulation of the PPN in vivo induced fast oscillation (20 - 40 Hz) in the cortical EEG3, while bilateral PPN lesions in the rat reduced or eliminated REM sleep4. While a majority of PPN neurons fire action potentials at beta/gamma-band frequency (20 - 80 Hz), some neurons presented low rates of spontaneous firing (&#60; 10 Hz) 5. Furthermore, the PPN seems to be involved in other aspects of behavior such as motivation and attention6. Direct high frequency (40 - 60 Hz)7 electrical stimulation of PPN nucleus in decerebrate animals can promote locomotion. In recent years, deep brain stimulation (DBS) of PPN has been used to treat patients suffering from disorders involving gait deficits such as Parkinson&#39;s disease (PD)8.
Previous reports demonstrated that almost all PPN neurons can fire action potentials at gamma band frequency when depolarized using square current pulses9. Because of the drastic activation of voltage-gated potassium channels during square pulses depolarizations up to or under -25 mV. As a consequence, no robust gamma oscillations were observed after blocking action potentials generation using tetrodotoxin10. In an effort to bypass such a problem, 1 - 2 sec long depolarizing current ramps were used. Ramps gradually depolarized the membrane potential from resting values up to 0 mV, while partially inactivating voltage-gated potassium channels. Clear gamma band membrane oscillations were evident within the voltage dependence window of high threshold calcium channels (i.e., between -25 mV and -0 mV) 10. In conclusion, gamma band activity was observed in PPN neurons9, and both P/Q- and N-type voltage-gated calcium channels need to be activated in order to generate gamma band oscillations in the PPN10.
A series of studies determined the location of high threshold calcium channels in PPN neurons. Injecting the combination of dyes, ratiometric fluorescence imaging showed calcium transients through voltage-gated calcium channels that are activated in different dendrites when depolarized using current ramps11.
Intrinsic properties of PPN neurons have been suggested to allow simultaneous activation of these cells during waking and REM sleep, thus inducing high-frequency oscillatory neuronal activity between the RAS and thalamocortical loops. Such long-reaching interaction is considered to support a brain state capable of reliably assessing the world around us on a continuous basis12. Here, we describe the experimental conditions necessary to generate and maintain gamma band oscillation in PPN cells in vitro. This protocol has not been described previously, and would help a number of groups to study intrinsic membrane properties mediating gamma band activity at other brain areas. Moreover, current steps might lead to the erroneous conclusion that gamma band activity cannot be generated in these cells.

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	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:619px;">Sucrose</td>
			<td style="width:335px;">Sigma-Aldrich</td>
			<td style="width:307px;">S8501</td>
			<td style="width:405px;">C12H22O11, molecular weight = 342.30</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sodium Bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S6014</td>
			<td>NaHCO3, molecular weight = 84.01</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Potassium Chloride</td>
			<td>Sigma-Aldrich</td>
			<td>P3911</td>
			<td>KCl, molecular weight = 74.55</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Magnesium Chloride Hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>M9272</td>
			<td>MgCl2 &#183; 6H2O, molecular weight =&#160; 203.30</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Calcium Chloride Dihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>C3881</td>
			<td>CaCl2 &#183; 2H2O, molecular weight =147.02</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">D-(+)-Glucose</td>
			<td>Sigma-Aldrich</td>
			<td>G5767</td>
			<td>C6H12O6, molecular weight = 180.16</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">L-Ascorbic Acid</td>
			<td>Sigma-Aldrich</td>
			<td>A5960</td>
			<td>C6H8O6, molecular weight =176.12</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sodium Chloride</td>
			<td>Acros Organics</td>
			<td>327300025</td>
			<td>NaCl, molecular weight =&#160; 58.44</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Potassium Gluconate</td>
			<td>Sigma-Aldrich</td>
			<td>G4500</td>
			<td>C6H11KO7, molecular weight =&#160; 234.25</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Phosphocreatine di(tris) salt</td>
			<td>Sigma-Aldrich</td>
			<td>P1937</td>
			<td>C4H10N3O5P &#183; 2C4H11NO3, molecular weight =&#160; 453.38</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375</td>
			<td>C8H18N2O4S, molecular weight = 238.30</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">EGTA</td>
			<td>Sigma-Aldrich</td>
			<td>E0396</td>
			<td>[-CH2OCH2CH2N(CH2CO2H)2]2, molecular weight = 380.40</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Adenosine 5&#39;-triphosphate magnesium salt</td>
			<td>Sigma-Aldrich</td>
			<td>A9187</td>
			<td>&#160;C10H16N5O13P3 &#183; xMg2+, molecular weight = 507.18</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Guanosine 5&#39;-triphosphate sodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>G8877</td>
			<td>C10H16N5O14P3 &#183; xNa+, molecular weight = 523.18</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Tetrodotoxin citrate</td>
			<td>Alomone Labs</td>
			<td>T-550</td>
			<td>C11H17N3O8, molecular weight = 319.27</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">&#160;DL-2-Amino-5-Phosphonovaleric Acid</td>
			<td>Sigma-Aldrich</td>
			<td>A5282</td>
			<td>&#160;C5H12NO5P, molecular weight = 197.13</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">CNQX disodium salt hydrate&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>C239</td>
			<td>C9H2N4Na2O4 &#183; xH2O, molecular weight = 276.12</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Strychnine</td>
			<td>Sigma-Aldrich</td>
			<td>S0532</td>
			<td>C21H22N2O2, molecular weight = 334.41</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Mecamylamine hydrochloride</td>
			<td>Sigma-Aldrich</td>
			<td>M9020</td>
			<td>&#160;C11H21N &#183; HCl, molecular weight = 203.75</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Gabazine (SR-95531)</td>
			<td>Sigma-Aldrich</td>
			<td>S106</td>
			<td>C15H18BrN3O3, molecular weight = 368.23</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Ketamine hydrochloride</td>
			<td>Mylan</td>
			<td>67457-001-00</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Microscope</td>
			<td>Nikon</td>
			<td>Eclipse E600FN</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>ROE-200</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>MPC-200</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Amplifier</td>
			<td>Molecular Devices</td>
			<td>Multiclamp 700B</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">A/D converter</td>
			<td>Molecular Devices</td>
			<td>Digidata 1440A</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Heater</td>
			<td>Warner Instruments</td>
			<td>TC-324B</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Pump</td>
			<td>Cole-Parmer</td>
			<td>Masterflex L/S 7519-20</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Pump cartridge</td>
			<td>Cole-Parmer</td>
			<td>Masterflex 7519-85</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Pipette puller</td>
			<td>Sutter Instruments</td>
			<td>P-97</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Camera</td>
			<td>Q-Imaging</td>
			<td>RET-200R-F-M-12-C</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Vibratome</td>
			<td>Leica Biosystems</td>
			<td>&#160;Leica VT1200 S</td>
			<td></td>
		</tr>
		<tr height="33">
			<td height="33" style="height:33px;">Refrigeration system</td>
			<td>Vibratome Instruments</td>
			<td>900R</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">Equipment</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">microscope</td>
			<td>Nikon</td>
			<td>Eclipse E600FN</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>ROE-200</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>MPC-200</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">amplifier</td>
			<td>Molecular Devices</td>
			<td>Multiclamp 700B</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">A/D converter</td>
			<td>Molecular Devices</td>
			<td>Digidata 1440A</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">heater</td>
			<td>Warner Instruments</td>
			<td>TC-324B</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">pump</td>
			<td>Cole-Parmer</td>
			<td>Masterflex L/S 7519-20</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">pump cartridge</td>
			<td>Cole-Parmer</td>
			<td>Masterflex 7519-85</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">pipette puller</td>
			<td>Sutter Instruments</td>
			<td>P-97</td>
			<td></td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;">camera</td>
			<td>Q-Imaging</td>
			<td>RET-200R-F-M-12-C</td>
			<td></td>
		</tr>
	</tbody>
</table>

recording gamma band oscillations in pedunculopontine nucleus neurons

Synaptic efferents from the PPN are known to modulate the neuronal activity of several intralaminar thalamic regions (e.g., the centrolateral/parafascicular; Cl/Pf nucleus). The activation of either the PPN or Cl/Pf nuclei in vivo has been described to induce the arousal of the animal and an increment in gamma band activity in the cortical electroencephalogram (EEG). The cellular mechanisms for the generation of gamma band oscillations in Reticular Activating System (RAS) neurons are the same as those found to generate gamma band oscillations in other brains nuclei. During current-clamp recordings of PPN neurons (from parasagittal slices from 9 - 25 day-old rats), the use of depolarizing square steps rapidly activated voltage-dependent potassium channels that prevented PPN neurons from being depolarized beyond -25 mV.
Injecting 1 - 2 sec long depolarizing current ramps gradually depolarized PPN membrane potential resting values towards 0 mV. However, injecting depolarizing square pulses generated gamma-band oscillations of membrane potential that showed to be smaller in amplitude compared to the oscillations generated by ramps. All experiments were performed in the presence of voltage-gated sodium channels and fast synaptic receptors blockers. It has been shown that the activation of high-threshold voltage-dependent calcium channels underlie gamma-band oscillatory activity in PPN neurons. Specific methodological and pharmacological interventions are described here, providing the necessary tools to induce and sustain PPN subthreshold gamma band oscillation in vitro.

Initially, gamma oscillations were evoked using square current pulses. Current clamp recording of PPN neurons in the presence of synaptic blockers and TTX was continuously monitored to assure that resting membrane potential was kept stable at ~-50 mV (Figure 1A). Two second long square current pulses were injected intracellularly by the patch clamp amplifier through the recording pipette, increasing their amplitude from 200 pA to 600 pA (Figure 1A). Membr...

Watch this Scientific Journal Video about Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons at JoVE.com

Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons

The pedunculopontine nucleus (PPN) is located in the brainstem and its neurons are maximally activated during waking and rapid eye movement (REM) sleep brain states. This work describes the experimental approach to record in vitro gamma band subthreshold membrane oscillation in PPN neurons.

Synaptic efferents from the PPN are known to modulate the neuronal activity of several intralaminar thalamic regions (e.g., the ...

recording-gamma-band-oscillations-in-pedunculopontine-nucleus-neurons

Research

JoVE Journal

Neuroscience

8.6K Views.  University of Buenos Aires. The pedunculopontine nucleus (PPN) is located in the brainstem and its neurons are maximally activated during waking and rapid eye movement (REM) sleep brain states. This work describes the experimental approach to record in vitro gamma band subthreshold membrane oscillation in PPN neurons.

Video: Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons

Slice Preparation, Organotypic Tissue Culturing and Luciferase Recording of Clock Gene Activity in the Suprachiasmatic Nucleus

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the anterior hypothalamus. The clock is directly synchronized by light via the retina and optic nerve. Circadian oscillations are generated by interacting negative feedback loops of a number of so called "clock genes" and their protein products, including the Period (Per) genes. The core clock is also dependent on membrane depolarization, calcium and cAMP 1. The SCN shows daily oscillations in clock gene expression, metabolic activity and spontaneous electrical activity. Remarkably, this endogenous cyclic activity persists in adult tissue slices of the SCN 2-4. In this way, the biological clock can easily be studied in vitro, allowing molecular, electrophysiological and metabolic investigations of the pacemaker function.
The SCN is a small, well-defined bilateral structure located right above the optic chiasm 5. In the rat it contains ~8.000 neurons in each nucleus and has dimensions of approximately 947 &mu;m (length, rostrocaudal axis) x 424 &mu;m (width) x 390 &mu;m (height) 6. To dissect out the SCN it is necessary to cut a brain slice at the specific level of the brain where the SCN can be identified. Here, we describe the dissecting and slicing procedure of the SCN, which is similar for mouse and rat brains. Further, we show how to culture the dissected tissue organotypically on a membrane 7, a technique developed for SCN tissue culture by Yamazaki et al. 8. Finally, we demonstrate how transgenic tissue can be used for measuring expression of clock genes/proteins using dynamic luciferase reporter technology, a method that originally was used for circadian measurements by Geusz et al. 9. We here use SCN tissues from the transgenic knock-in PERIOD2::LUCIFERASE mice produced by Yoo et al. 10. The mice contain a fusion protein of PERIOD (PER) 2 and the firefly enzyme LUCIFERASE. When PER2 is translated in the presence of the substrate for luciferase, i.e. luciferin, the PER2 expression can be monitored as bioluminescence when luciferase catalyzes the oxidation of luciferin. The number of emitted photons positively correlates to the amount of produced PER2 protein, and the bioluminescence rhythms match the PER2 protein rhythm in vivo 10. In this way the cyclic variation in PER2 expression can be continuously monitored real time during many days. The protocol we follow for tissue culturing and real-time bioluminescence recording has been thoroughly described by Yamazaki and Takahashi 11.

The procedure of preparing slices containing the adult mouse hypothalamic suprachiasmatic nucleus (SCN), and a rapid way to culture the SCN tissue in organotypic culture condition, are reported. Further, the measurement of oscillatory clock gene protein expression using dynamic luciferase reporter technology is described.

A central circadian (~24 hr) clock coordinating daily rhythms in physiology and behavior resides in the suprachiasmatic nucleus (SCN) located in the ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of neurophysiology, neuroethology, and developmental biology. The goal of this report is to consolidate experimental techniques from the leech system into a single article that will be of use to physiologists with expertise in other nervous system preparations, or to biology students with little or no electrophysiology experience. We demonstrate how to dissect the leech for recording intracellularly from identified neural circuits in the ganglion. Next we show how individual cells of known function can be removed from the ganglion to be cultured in a Petri dish, and how to record from those neurons in culture. Then we demonstrate how to prepare a patch of innervated skin to be used for mapping sensory or motor fields. These leech preparations are still widely used to address basic electrical properties of neural networks, behavior, synaptogenesis, and development. They are also an appropriate training module for neuroscience or physiology teaching laboratories.

Intracellular Recording, Sensory Field Mapping, and Culturing Identified Neurons in the Leech, Hirudo medicinalis

This article describes three nervous system preparations using leeches: intracellular recording from neurons in ventral ganglia, culturing neurons from ventral ganglia, and recording from a patch of innervated skin to map sensory fields.

The freshwater leech, Hirudo medicinalis, is a versatile model organism that has been used to address scientific questions in the fields of ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Perforated Patch-clamp Recording of Mouse Olfactory Sensory Neurons in Intact Neuroepithelium: Functional Analysis of Neurons Expressing an Identified Odorant Receptor

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of olfactory-driven behaviors and their modulation. These coding properties depend heavily on the initial interaction between odor molecules and the olfactory receptor (OR) expressed in the OSNs. The identity, specificity and ligand spectrum of the expressed OR are critical. The probability to find the ligand of the OR expressed in an OSN chosen randomly within the epithelium is very low. To address this challenge, this protocol uses genetically tagged mice expressing the fluorescent protein GFP under the control of the promoter of defined ORs. OSNs are located in a tight and organized epithelium lining the nasal cavity, with neighboring cells influencing their maturation and function. Here we describe a method to isolate an intact olfactory epithelium and record through patch-clamp recordings the properties of OSNs expressing defined odorant receptors. The protocol allows one to characterize OSN membrane properties while keeping the influence of the neighboring tissue. Analysis of patch-clamp results yields&#160;a precise quantification of ligand/OR interactions, transduction pathways and pharmacology, OSNs&#39; coding properties and their modulation at the membrane level.&#160;

Analyzing the physiological properties of olfactory sensory neurons still faces technical limitations. Here we record them through perforated patch-clamp in an intact preparation of the olfactory epithelium in gene-targeted mice. This technique allows the characterization of membrane properties and responses to specific ligands of neurons expressing defined olfactory receptors.

Analyzing the physiological responses of olfactory sensory neurons (OSN) when stimulated with specific ligands is critical to understand the basis of ...

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent improvements in methodologies and applications for the study of theta oscillations. A simple characterization of the isolated hippocampal preparation is presented whereby the relationship between internal hippocampal theta oscillators is examined together with the activity of pyramidal cells, and GABAergic interneurons, of the cornu ammonis-1 (CA1) and subiculum (SUB) areas. Overall, we show that the isolated hippocampus is capable of generating intrinsic theta oscillations in vitro and that rhythmicity generated within the hippocampus can be precisely manipulated by optogenetic stimulation of parvalbumin-positive (PV) interneurons. The in vitro isolated hippocampal preparation offers a unique opportunity to use simultaneous field and intracellular patch-clamp recordings from visually-identified neurons to better understand the mechanisms underlying theta rhythm generation.

Tuning in the Hippocampal Theta Band In Vitro: Methodologies for Recording from the Isolated Rodent Septohippocampal Circuit

Here, we present a protocol for recording rhythmic neuronal network theta and gamma oscillations from an isolated whole hippocampal preparation. We describe the experimental steps from extraction of the hippocampus to details of field, unitary and whole-cell patch clamp recordings as well as optogenetic pacing of the theta rhythm.

This protocol outlines the procedures for preparing and recording from the isolated whole hippocampus, of WT and transgenic mice, along with recent ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in regulating neuronal growth and development. However, the mechanisms underlying Drd1&#945; receptor abnormalities mediating behavioral responses and modulating working memory function are still unclear. Using a novel RNA in situ hybridization assay, the current study identified dopamine Drd1&#945; receptor and tyrosine hydroxylase (TH) RNA expression from DA-related circuitry in the nucleus accumbens (NAc) area and substantia nigra region (SNR), respectively. Drd1&#945; expression in the NAc shows a &#34;discrete dot&#34; staining pattern. Clear sex differences in Drd1&#945; expression were observed. In contrast, TH shows a &#34;clustered&#34; staining pattern. Regarding TH expression, female rats displayed a higher signal expression per cell relative to male animals. The methods presented here provide a novel in situ hybridization technique for investigating changes in dopamine system dysfunction during the progression of central nervous system diseases.

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

Identification of dopamine D1-alpha receptor in the nucleus accumbens is critical for clarifying D1 receptor dysfunction during a central nervous system disease. We performed a novel RNA in situ hybridization assay to visualize single RNA molecules in a specific brain area.