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 cortical gamma activity on the EEG2,3,5,18-21. Gamma band neural activity has been reported in the mouse PPN in vitro22, in the rat REM sleep-induction area (to which the PPN projects23), in the cat in vivo3, in the PPN area in primates when locomoting24, and in the region of the PPN in humans during stepping25. That is, there is ample evidence for gamma band activity in the PPN across species.
This experimental protocol detailed here permitted the description of gamma oscillations present at all rat PPN cell types 10. In fact, intrinsic mechanisms underlying gamma oscillations were described to be present in every PPN neuron (while cells around the PPN do not share that property), regardless of previous used classifications or synaptic transmitter type: type I, II, or III, or transmitter type, cholinergic, glutamatergic or GABAergic share the same gamma-band generating property10. The use of current ramps has the limitation of requiring continuous monitoring of the resting membrane potential, and permanent bridge compensation throughout the experiment. In some cases, ramps shorter than 1 sec were observed to not fully activate voltage-gated calcium channels, while ramp durations of 5 sec or longer resulted in membrane resistance changes that would remain uncompensated during the experiment. In case no gamma oscillations were observed, small adjustments in ramp duration could enhance gamma band oscillation amplitudes.
Combining depolarizing current-ramps with specific toxins we have described that in an important proportion of PPN cells (~50%) blocking N-and P/Q-type (using both &#969;-CgTx and &#969;-Aga) channels calcium channels reduced gamma oscillation amplitude, which allow us to classified them as P/Q- and N-type cells. In other cells (20%), gamma oscillations were only affected by &#969;-Aga, suggesting that these cells expressed only P/Q-type channels. In the rest of the cells (30%) only &#969;-CgTx blocked them, suggesting these PPN neurons had only N-type channels. These results confirmed the presence of cells in the PPN that manifest gamma band oscillations through the expression of different voltage-gated calcium channels 26,27. This new protocol can also be considered a key tool that allowed the introduction of a new PPN cell type classification to the field. In fact, it has been suggested that N-type only PPN neurons fire only during REM sleep (&#34;REM-on&#34;), P/Q-type only during waking (&#34;Wake-on&#34;), or N-type + P/Q-type during both waking and REM sleep (&#34;Wake/REM-on&#34;) 26,27. Furthermore, this protocol might be used in future experiments to describe oscillatory activity at other brain areas and to demonstrate the intrinsic properties triggering them.

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.

<table><tbody><tr><td>Sucrose</td><td>Sigma-Aldrich</td><td>S8501</td><td>C&lt;sub&gt;12&lt;/sub&gt;H&lt;sub&gt;22&lt;/sub&gt;O&lt;sub&gt;11&lt;/sub&gt;, molecular weight = 342.30</td></tr><tr><td>Sodium Bicarbonate</td><td>Sigma-Aldrich</td><td>S6014</td><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;, molecular weight = 84.01</td></tr><tr><td>Potassium Chloride</td><td>Sigma-Aldrich</td><td>P3911</td><td>KCl, molecular weight = 74.55</td></tr><tr><td>Magnesium Chloride Hexahydrate</td><td>Sigma-Aldrich</td><td>M9272</td><td>MgCl&lt;sub&gt;2&lt;/sub&gt; &amp;middot; 6H&lt;sub&gt;2&lt;/sub&gt;O, molecular weight =&amp;nbsp; 203.30</td></tr><tr><td>Calcium Chloride Dihydrate</td><td>Sigma-Aldrich</td><td>C3881</td><td>CaCl&lt;sub&gt;2&lt;/sub&gt; &amp;middot; 2H&lt;sub&gt;2&lt;/sub&gt;O, molecular weight =147.02</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma-Aldrich</td><td>G5767</td><td>C&lt;sub&gt;6&lt;/sub&gt;H&lt;sub&gt;12&lt;/sub&gt;O&lt;sub&gt;6&lt;/sub&gt;, molecular weight = 180.16</td></tr><tr><td>L-Ascorbic Acid</td><td>Sigma-Aldrich</td><td>A5960</td><td>C&lt;sub&gt;6&lt;/sub&gt;H&lt;sub&gt;8&lt;/sub&gt;O&lt;sub&gt;6&lt;/sub&gt;, molecular weight =176.12</td></tr><tr><td>Sodium Chloride</td><td>Acros Organics</td><td>327300025</td><td>NaCl, molecular weight =&amp;nbsp; 58.44</td></tr><tr><td>Potassium Gluconate</td><td>Sigma-Aldrich</td><td>G4500</td><td>C&lt;sub&gt;6&lt;/sub&gt;H&lt;sub&gt;11&lt;/sub&gt;KO&lt;sub&gt;7&lt;/sub&gt;, molecular weight =&amp;nbsp; 234.25</td></tr><tr><td>Phosphocreatine di(tris) salt</td><td>Sigma-Aldrich</td><td>P1937</td><td>C&lt;sub&gt;4&lt;/sub&gt;H&lt;sub&gt;10&lt;/sub&gt;N&lt;sub&gt;3&lt;/sub&gt;O&lt;sub&gt;5&lt;/sub&gt;P &amp;middot; 2C&lt;sub&gt;4&lt;/sub&gt;H&lt;sub&gt;11&lt;/sub&gt;NO&lt;sub&gt;3&lt;/sub&gt;, molecular weight =&amp;nbsp; 453.38</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich</td><td>H3375</td><td>C&lt;sub&gt;8&lt;/sub&gt;H&lt;sub&gt;18&lt;/sub&gt;N&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;4&lt;/sub&gt;S, molecular weight = 238.30</td></tr><tr><td>EGTA</td><td>Sigma-Aldrich</td><td>E0396</td><td>[-CH&lt;sub&gt;2&lt;/sub&gt;OCH&lt;sub&gt;2&lt;/sub&gt;CH&lt;sub&gt;2&lt;/sub&gt;N(CH&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;2&lt;/sub&gt;H)&lt;sub&gt;2&lt;/sub&gt;]&lt;sub&gt;2&lt;/sub&gt;, molecular weight = 380.40</td></tr><tr><td>Adenosine 5&#039;-triphosphate magnesium salt</td><td>Sigma-Aldrich</td><td>A9187</td><td>&amp;nbsp;C&lt;sub&gt;10&lt;/sub&gt;H&lt;sub&gt;16&lt;/sub&gt;N&lt;sub&gt;5&lt;/sub&gt;O&lt;sub&gt;13&lt;/sub&gt;P&lt;sub&gt;3&lt;/sub&gt; &amp;middot; xMg2+, molecular weight = 507.18</td></tr><tr><td>Guanosine 5&#039;-triphosphate sodium salt hydrate</td><td>Sigma-Aldrich</td><td>G8877</td><td>C&lt;sub&gt;10&lt;/sub&gt;H&lt;sub&gt;16&lt;/sub&gt;N&lt;sub&gt;5&lt;/sub&gt;O&lt;sub&gt;14&lt;/sub&gt;P&lt;sub&gt;3&lt;/sub&gt; &amp;middot; xNa+, molecular weight = 523.18</td></tr><tr><td>Tetrodotoxin citrate</td><td>Alomone Labs</td><td>T-550</td><td>C&lt;sub&gt;11&lt;/sub&gt;H&lt;sub&gt;17&lt;/sub&gt;N&lt;sub&gt;3&lt;/sub&gt;O&lt;sub&gt;8&lt;/sub&gt;, molecular weight = 319.27</td></tr><tr><td>&amp;nbsp;DL-2-Amino-5-Phosphonovaleric Acid</td><td>Sigma-Aldrich</td><td>A5282</td><td>&amp;nbsp;C&lt;sub&gt;5&lt;/sub&gt;H&lt;sub&gt;12&lt;/sub&gt;NO&lt;sub&gt;5&lt;/sub&gt;P, molecular weight = 197.13</td></tr><tr><td>CNQX disodium salt hydrate&amp;nbsp;</td><td>Sigma-Aldrich</td><td>C239</td><td>C&lt;sub&gt;9&lt;/sub&gt;H&lt;sub&gt;2&lt;/sub&gt;N&lt;sub&gt;4&lt;/sub&gt;Na&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;4&lt;/sub&gt; &amp;middot; xH2O, molecular weight = 276.12</td></tr><tr><td>Strychnine</td><td>Sigma-Aldrich</td><td>S0532</td><td>C&lt;sub&gt;21&lt;/sub&gt;H&lt;sub&gt;22&lt;/sub&gt;N&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;2&lt;/sub&gt;, molecular weight = 334.41</td></tr><tr><td>Mecamylamine hydrochloride</td><td>Sigma-Aldrich</td><td>M9020</td><td>&amp;nbsp;C&lt;sub&gt;11&lt;/sub&gt;H&lt;sub&gt;21&lt;/sub&gt;N &amp;middot; HCl, molecular weight = 203.75</td></tr><tr><td>Gabazine (SR-95531)</td><td>Sigma-Aldrich</td><td>S106</td><td>C&lt;sub&gt;15&lt;/sub&gt;H&lt;sub&gt;18&lt;/sub&gt;BrN&lt;sub&gt;3&lt;/sub&gt;O&lt;sub&gt;3&lt;/sub&gt;, molecular weight = 368.23</td></tr><tr><td>Ketamine hydrochloride</td><td>Mylan</td><td>67457-001-00</td><td></td></tr><tr><td>Microscope</td><td>Nikon</td><td>Eclipse E600FN</td><td></td></tr><tr><td>Micromanipulator</td><td>Sutter Instruments</td><td>ROE-200</td><td></td></tr><tr><td>Micromanipulator</td><td>Sutter Instruments</td><td>MPC-200</td><td></td></tr><tr><td>Amplifier</td><td>Molecular Devices</td><td>Multiclamp 700B</td><td></td></tr><tr><td>A/D converter</td><td>Molecular Devices</td><td>Digidata 1440A</td><td></td></tr><tr><td>Heater</td><td>Warner Instruments</td><td>TC-324B</td><td></td></tr><tr><td>Pump</td><td>Cole-Parmer</td><td>Masterflex L/S 7519-20</td><td></td></tr><tr><td>Pump cartridge</td><td>Cole-Parmer</td><td>Masterflex 7519-85</td><td></td></tr><tr><td>Pipette puller</td><td>Sutter Instruments</td><td>P-97</td><td></td></tr><tr><td>Camera</td><td>Q-Imaging</td><td>RET-200R-F-M-12-C</td><td></td></tr><tr><td>Vibratome</td><td>Leica Biosystems</td><td>&amp;nbsp;Leica VT1200 S</td><td></td></tr><tr><td>Refrigeration system</td><td>Vibratome Instruments</td><td>900R</td><td></td></tr><tr><td>&lt;strong&gt;Equipment&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>microscope</td><td>Nikon</td><td>Eclipse E600FN</td><td></td></tr><tr><td>micromanipulator</td><td>Sutter Instruments</td><td>ROE-200</td><td></td></tr><tr><td>micromanipulator</td><td>Sutter Instruments</td><td>MPC-200</td><td></td></tr><tr><td>amplifier</td><td>Molecular Devices</td><td>Multiclamp 700B</td><td></td></tr><tr><td>A/D converter</td><td>Molecular Devices</td><td>Digidata 1440A</td><td></td></tr><tr><td>heater</td><td>Warner Instruments</td><td>TC-324B</td><td></td></tr><tr><td>pump</td><td>Cole-Parmer</td><td>Masterflex L/S 7519-20</td><td></td></tr><tr><td>pump cartridge</td><td>Cole-Parmer</td><td>Masterflex 7519-85</td><td></td></tr><tr><td>pipette puller</td><td>Sutter Instruments</td><td>P-97</td><td></td></tr><tr><td>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). Membrane potential was depolarized to voltage levels closer to -20 mV when 600 pA, resulting in small amplitude gamma oscillations. Powers spectra of gamma oscillations presented very small amplitude values (i.e., &#60; 0.01; Figure 1B). On the other hand, two second depolarizing current ramps was injected intracellularly by the patch clamp amplifier through the recording pipette, generating robust gamma oscillations (Figure 2A) that presented much higher power amplitudes (Figure 2B).
Previous work10 has linked differences between square and ramp current protocols to the string activation of voltage-gated potassium channels during sudden depolarization of the former. Slow depolarizing ramps might instead be inactivating potassium channels.
<img alt="Figure 1" src="/files/ftp_upload/54685/54685fig1.jpg" /> 
Figure 1. Membrane Oscillations in Whole-cell Recorded PPN Neurons using Current Square Pulses of Increasing Amplitude. (A) Representative membrane potential responses to depolarizing 2 sec long square steps of increasing amplitude injected intracellularly by the patch clamp amplifier through the recording pipette, in the presence of synaptic blockers and TTX. (B) Overlapping power spectra amplitudes for oscillations obtained using square pulses shown in A. Power spectra were obtained using a Hamming window function after 20 - 60 Hz bandpass filtering oscillations generated by the depolarizing steps. PPN neuron was recorded in current-clamp configuration combining high-K+ intracellular solution and synaptic blockers + TTX described in Protocol. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54685/54685fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54685/54685fig2.jpg" /> 
Figure 2. Gamma Band Membrane Oscillations in Whole-cell Recorded PPN Neurons using Current Ramps of Augmenting Amplitude. (A) Representative membrane potential responses to depolarizing 2 sec long ramps injected by the patch clamp amplifier intracellularly through the recording pipette, showing increasing amplitude in the presence of synaptic blockers and TTX. (B) Overlapping power spectra amplitudes for oscillations obtained using ramps shown in A. Power spectra were obtained using a Hamming window function after 20 - 60 Hz bandpass filtering oscillations generated by the depolarizing ramp. PPN neuron was recorded in current-clamp configuration combining high-K+ intracellular solution and synaptic blockers + TTX described in Protocol. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54685/54685fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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

Universidad San Sebastian

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

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2 or record extracellular neuronal activity during a movement.3,4 While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo movement thus allowing substantial insight into the role of the neuron in sensorimotor function.

A technique is described to quantify the in vivo physiological response of mammalian neurons during movement and correlate the physiology of the neuron with neuronal morphology, neurochemical phenotype and synaptic microcircuitry.

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor ...

Automatic Detection of Highly Organized Theta Oscillations in the Murine EEG

Theta activity is generated in the septohippocampal system and can be recorded using deep intrahippocampal electrodes and implantable electroencephalography (EEG) radiotelemetry or tether system approaches. Pharmacologically, hippocampal theta is heterogeneous (see dualistic theory) and can be differentiated into type I and type II theta. These individual EEG subtypes are related to specific cognitive and behavioral states, such as arousal, exploration, learning and memory, higher integrative functions, etc. In neurodegenerative diseases such as Alzheimer&#39;s, structural and functional alterations of the septohippocampal system can result in impaired theta activity/oscillations. A standard quantitative analysis of the hippocampal EEG includes a Fast-Fourier-Transformation (FFT)-based frequency analysis. However, this procedure does not provide details about theta activity in general and highly-organized theta oscillations in particular. In order to obtain detailed information on highly-organized theta oscillations in the hippocampus, we have developed a new analytical approach. This approach allows for time- and cost-effective quantification of the duration of highly-organized theta oscillations and their frequency characteristics.

Theta activity in the hippocampus is related to specific cognitive and behavioral stages. Here, we describe an analytical method to detect highly-organized theta oscillations within the hippocampus using a time-frequency (i.e., wavelet analysis)-based approach.

Theta activity is generated in the septohippocampal system and can be recorded using deep intrahippocampal electrodes and implantable ...

Behavior

Manufacturing and Using Piggy-back Multibarrel Electrodes for In vivo Pharmacological Manipulations of Neural Responses

In vivo recordings from single neurons allow an investigator to examine the firing properties of neurons, for example in response to sensory stimuli. Neurons typically receive multiple excitatory and inhibitory afferent and/or efferent inputs that integrate with each other, and the ultimate measured response properties of the neuron are driven by the neural integrations of these inputs. To study information processing in neural systems, it is necessary to understand the various inputs to a neuron or neural system, and the specific properties of these inputs. A powerful and technically relatively simple method to assess the functional role of certain inputs that a given neuron is receiving is to dynamically and reversibly suppress or eliminate these inputs, and measure the changes in the neuron's output caused by this manipulation. This can be accomplished by pharmacologically altering the neuron's immediate environment with piggy-back multibarrel electrodes. These electrodes consist of a single barrel recording electrode and a multibarrel drug electrode that can carry up to 4 different synaptic agonists or antagonists. The pharmacological agents can be applied iontophoretically at desired times during the experiment, allowing for time-controlled delivery and reversible reconfiguration of synaptic inputs. As such, pharmacological manipulation of the microenvironment represents a powerful and unparalleled method to test specific hypotheses about neural circuit function.
Here we describe how piggy-back electrodes are manufactured, and how they are used during in vivo experiments. The piggy-back system allows an investigator to combine a single barrel recording electrode of any arbitrary property (resistance, tip size, shape etc) with a multibarrel drug electrode. This is a major advantage over standard multi-electrodes, where all barrels have more or less similar shapes and properties. Multibarrel electrodes were first introduced over 40 years ago 1-3, and have undergone a number of design improvements 2,3 until the piggy-back type was introduced in the 1980s 4,5. Here we present a set of important improvements in the laboratory production of piggy-back electrodes that allow for deep brain penetration in intact in vivo animal preparations due to a relatively thin electrode shaft that causes minimal damage. Furthermore these electrodes are characterized by low noise recordings, and have low resistance drug barrels for very effective iontophoresis of the desired pharmacological agents.

Manufacturing and Using Piggy-back Multibarrel Electrodes for In vivo Pharmacological Manipulations of Neural Responses

Iontophoresis of neural agonists and antagonists during extracellular in vivo recordings is a powerful way to manipulate a neuron&#x2019;s microenvironment. These manipulations can most easily be done via piggy-back multibarrel electrodes. Here we describe how to manufacture them and use them during auditory recordings.

In vivo recordings from single neurons allow an investigator to examine the firing properties of neurons, for example in response to sensory stimuli. ...

Generation of Local CA1 &#947; Oscillations by Tetanic Stimulation

Neuronal network oscillations are important features of brain activity in health and disease and can be modulated by a range of clinically used drugs. A protocol is provided to generate a model for studying CA1 &#947; oscillations (20 - 80 Hz). These &#947; oscillations are stable for at least 30 min&#160;and depend upon excitatory and inhibitory synaptic activity in addition to activation of pacemaker currents. Tetanically stimulated oscillations have a number of reproducible and easily quantifiable characteristics including spike count, oscillation duration, latency and frequency that report upon the network state. The advantages of the electrically stimulated oscillations include stability, reproducibility and episodic acquisition enabling robust characterization of network function. This model of CA1 &#947; oscillations can be used to study cellular mechanisms and to systematically investigate how neuronal network activity is altered in disease and by drugs. Disease state pharmacology can be readily incorporated by the use of brain slices from genetically modified or interventional animal models to enable selection of drugs that specifically target disease mechanisms.

Oscillations are fundamental network properties and are modulated by disease and drugs. Studying brain-slice oscillations allows characterization of isolated networks under controlled conditions. Protocols are provided for the preparation of acute brain slices for evoking CA1 &#947; oscillations.

Neuronal network oscillations are important features of brain activity in health and disease and can be modulated by a range of clinically used drugs. A ...

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 ...

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the pathophysiological basis of these diseases, it is necessary to precisely characterize in which way the processing of information is disturbed between the different neuronal parts of the circuit. Using extracellular in vivo electrophysiological recordings, it is possible to accurately delineate neuronal activity within a neuronal network. The application of this method has several advantages over alternative techniques, e.g., functional magnetic resonance imaging and calcium imaging, as it allows a unique temporal and spatial resolution and does not rely on genetically engineered organisms. However, the use of extracellular in vivo recordings is limited since it is an invasive technique that cannot be universally applied. In this article, a simple and easy to use method is presented with which it is possible to simultaneously record extracellular potentials such as local field potentials and multiunit activity at multiple sites of a network. It is detailed how a precise targeting of subcortical nuclei can be achieved using a combination of stereotactic surgery and online analysis of multi-unit recordings. Thus, it is demonstrated, how a complete network such as the hyperdirect cortico-basal ganglia loop can be studied in anesthetized animals in vivo.

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

In this study, the methodology is presented on how to perform multi-site in vivo electrophysiological recordings from the hyperdirect pathway under urethane anesthesia.

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the ...

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 ...

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ganglion cells and project it to the visual cortex. In addition, relay neurons receive top-down excitation from the cortex. The two main excitatory inputs to the relay neurons differ in several aspects. Each relay neuron receives input from only a few retinogeniculate synapses, which are large terminals with many release sites. This is reflected by the comparably strong excitation, the relay neurons receive, from retinal ganglion cells. Corticogeniculate synapses, in contrast, are simpler with few release sites and weaker synaptic strength. The two synapses also differ in their synaptic short-term plasticity. Retinogeniculate synapses have a high release probability and consequently display a short-term depression. In contrast, corticogeniculate synapses have a low release probability. Corticogeniculate fibers traverse the reticular thalamic nuclei before entering the lateral geniculate nucleus. The different locations of the reticular thalamic nucleus (rostrally from the lateral geniculate nucleus) and optic tract (ventro-laterally from the lateral geniculate nucleus) allow stimulating corticogeniculate or retinogeniculate synapses separately with extracellular stimulation electrodes. This makes the lateral geniculate nucleus an ideal brain area where two excitatory synapses with very different properties impinging onto the same cell type, can be studied simultaneously. Here, we describe a method to investigate the recording from relay neurons and to perform detailed analysis of the retinogeniculate and corticogeniculate synapse function in acute brain slices. The article contains a step-by-step protocol for the generation of acute brain slices of the lateral geniculate nucleus and steps for recording activity from relay neurons by stimulating the optic tract and corticogeniculate fibers separately.

Here, we present protocols for the preparation of acute brain slices containing the lateral geniculate nucleus and the electrophysiological investigation of retinogeniculate and corticogeniculate synapses function. This protocol provides an efficient way to study synapses with the high-release and low-release probability in the same acute brain slices.

The lateral geniculate nucleus is the first relay station for the visual information. Relay neurons of this thalamic nucleus integrate input from retinal ...

Recording Gamma Band Oscillations From Pedunculopontine Nucleus Neurons in a Brain Slice

Source: Urbano, F. J., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/54685">Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons.</a> J. Vis. Exp. (2016).
The video demonstrates the method to record gamma-band oscillations in PPN neurons. Using a sagittal brain section with synaptic blockers in aCSF, it employs positive current pulses via a patch pipette to open calcium channels, inducing crucial subthreshold activity for wakefulness.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Recording Gamma Band Oscillations in Pedunculopontine Nucleus Neurons

Summary

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Chapters in this video

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

Automatic Detection of Highly Organized Theta Oscillations in the Murine EEG

Manufacturing and Using Piggy-back Multibarrel Electrodes for In vivo Pharmacological Manipulations of Neural Responses

Generation of Local CA1 γ Oscillations by Tetanic Stimulation

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

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

Electrophysiological Investigations of Retinogeniculate and Corticogeniculate Synapse Function

Recording Gamma Band Oscillations From Pedunculopontine Nucleus Neurons in a Brain Slice