The authors would like to thank Isabelle Reus for establishment of tracing the electrode insertion side, Tobias Rosenbaum for LabView programming, Anneke Meyer for data analyzes and helpful discussions. We thank Randolf Menzel for discussion and practical help during early stage of electrode development. Furthermore we thank Brian Smith for postdoctoral association to MS-B. This work was supported by the Deutsche Forschungsgemeinschaft (DFG, SPP 1392, Ro1177/5-2) to WR.

1. Electrode Building (Figure 1)
<ol>
	<li>Production of an electrode adapter that fits the electrode interface board of commercial multi channel amplifier systems1,2,25.
	<ol>
		<li>Use a small Plexiglas plate glued to an 18 Pin connector base.</li>
		<li>Connect the base with 3&#160;short pieces of insulated wire to 3&#160;separate soldering lugs screwed onto the Plexiglas plate (Figure 1 A1-A3).</li>
		<li>Insert a groove into the Plexiglas plate in which a glass capillary can easily move and be held in place by a screw (Figure&#160;1 B1).</li>
		<li>Extend the glass capillary about 5 mm using a minutien pin.</li>
		<li>Attach the micro electrode wires along the minutien pin and the glass capillary to guarantee stabilization and support.</li>
	</ol>
	</li>
	<li>Multi-channel micro wire production (adopted from Ryuichy Okada32,33)
	<ol>
		<li>Span 3&#160;micro wires (polyurethane coated copper wire, 15 &#181;m diameter) in a way that they are placed next to each other (Figure&#160;1 B2).</li>
		<li>Use a 12 V soldering needle to spread a thin film of low melting dental wax (50 &#176;C) partially along the wires to glue them together (electrode tip) (Figure&#160;1 B3). Leave a few centimeters unglued (electrode end) as this section will be used later to connect the micro wires with the electrode adapter.</li>
	</ol>
	</li>
	<li>Connect the multi channel micro wire to the electrode adapter
	<ol>
		<li>Remove the glass capillary from the holder and attach it with the minutien pin to the electrode tip. Bring it into a position parallel to the micro-electrode (Figure1 B3).</li>
		<li>Glue the electrode tip to the minutien pin using low melting dental wax and cut the micro electrode at the tip, protruding 2-3 cm from the minutien pin and at the electrode end (small arrows Figure&#160;1 B3).</li>
		<li>Slip the capillary slightly back into the electrode adapter. Use the screw to fix it (Figure&#160;1 B4).</li>
		<li>Solder the loose ends of the three wires to the soldering lugs using a soldering gun with a temperature of around 360 &#176;C to ensure melting of the insulation (Figure&#160;1 B4). After soldering, ensure that there is adequate electrical contact (~300 kOhm).</li>
		<li>Mount one of the electrodes (master) to the electrode interface board of the headstage and fix the other multi channel electrode (slave) on a separate adapter. Connect the channels of the slave to the master electrode (Figure&#160;1 C1). Additionally, solder the reference as well as muscle electrodes to the master electrode base&#160;(Figure&#160;1 C2).</li>
	</ol>
	</li>
</ol>
2. Bee Preparation (Figure 2)
In these described experiments, the honeybee (Apis mellifera), which is an invertebrate animal and therefore does not require specific ethical permits for usage, is used.
<ol>
	<li>Catch honeybee foragers (A. mellifera) at the hive entrance in the morning as shown by others34,35.</li>
	<li>Chill the bees on crushed ice till immobilization (5 to 10 min) and fix one in a standard Plexiglas holder or metal tube in a way that the head is exposed (Figure&#160;2 A). For minimizing head movements use low-melting dental wax (~50 &#176;C) and fix the head carefully to the holder around the basis of the compound eyes and the neck.</li>
	<li>Use low melting wax to fixate the scapi of the antennae on the head capsule (Figure&#160;2 B) without touching the flagellum. The flagellum of the antenna needs to be pointed forward. Ensure that the bee can freely move its proboscis.</li>
	<li>Shave the head capsule to ensure an undisturbed view and access to the top of the head.</li>
	<li>Feed the bee with a 30% sucrose solution until saturation to ensure sufficient moistening of brain tissue and good viability of the animal (Figure 2 C).</li>
	<li>Make careful incisions vertically along the borders of the compound eyes and horizontally above the antennal bases as well as beneath the ocelli and remove the loosened piece of cuticle (Figure&#160;2 D).</li>
	<li>Carefully set aside hypopharyngeal glands and remove the trachea to ensure a clear view and access to the brain prior to electrode insertion (Figures 2E, 2F).</li>
</ol>
3. Electrode Insertion
In the example case illustrated in Figure &#160;2, one electrode is positioned aiming at the l-ALT, the other one aiming on the m-ALT2. Using particular landmarks, other target regions are possible as well, for example the AL and MB output regions1.
<ol>
	<li>Position the electrodes using micromanipulators at the region of interest (Figure&#160;2G and 3A). To target the m-ALT place the electrode between the AL and medially from the vertical lobe of the MB. Ensure the insertion site being above the branching point of the mediolateral ALT PNs. Protrude the electrode into the brain with a depth of about 180 &#181;m (Figure&#160;2E). For l-ALT PN recording place the electrode below the lateral protocerebrum (the LH) in the middle of an imaginary line between the lateral side of vertical lobe and the middle of the AL. Insert the electrode with a depth of about 300 &#181;m (Figure&#160;2E)</li>
	<li>Insert the reference (silver wire, about 25 &#181;m diameter) into the ipsilateral compound eye through a small cut in the cuticle. Insert another silver wire into the muscle projection region below the lateral ocelli. NOTE: If necessary the learning behavior of the bee can be monitored with high temporal precision by recording the muscle M17, which is involved in the proboscis extension response (PER) of the bee36 as described in25.</li>
	<li>To solidly anchor the electrodes within the brain and the head capsule, cover the entire space above the brain with two component silicon (Figure&#160;2H), which will prevent the brain from drying out. NOTE: the recordings can last for hours up to days, and the bees can for example be recorded during a classical conditioning procedure (Figure&#160;2H) or stimulated with a large panel of different odors.</li>
</ol>
4. Data Acquisition and Preprocessing
<ol>
	<li>Use proper acquisition software that meets the following requirements: sampling rate of min&#160;25 kHz; analogue1,25 or digital2 cross differentiation between the electrode channels; band pass filter from 300 Hz to 8,000 Hz to extract spike events.</li>
	<li>Use available spike sorting software to extract single unit activity, for example template matching techniques as included in the Spike2 software (Figure 3).</li>
	<li>For further analysis use the time stamps of the extracted units to compute single unit averaged odor responses (Figure 4) or to calculate population vectors for Principal Component Analysis (PCA) (Figure 5) using commercially available software. To further analyze single unit and population response latency please compare recent publications1,2,25.</li>
</ol>
5. Visualization of Relative Electrode Position (Figure 4)
<ol>
	<li>Dip the electrodes tips into a solution of either 5% Alexa&#160;hydrazide 568 or 5% Alexa hydrazide 488 which is dissolved in 0.5 M potassium chloride solution prior to the recording experiments.</li>
	<li>Remove the electrodes and the covering silicon carefully after the experiments, rinse the brain with bee Ringer solution, remove glands and trachea and insert tiny crystals of tetramethylrhodamin dextran or insert a 5% solution solved in 1.0 M potassium acetate into the AL to label the ALTs anterogradely. Perform the following steps in darkness.</li>
	<li>Allow the dye to be taken up and transported by the projection neurons along their axonal tracts (30-45 min) (Figure 4A), before washing the brain with bee Ringer&#160;solution three times for another 30-45 min.</li>
	<li>Chill the bee on ice until immobilization and carefully remove the brain from the head capsule. Fixate the brain by rinsing it in a 0.1 M PBS solution containing 4% formaldehyde and keep it overnight at 4 &#176;C.</li>
	<li>Wait at least 12 hr before washing the brain two times in 0.1 M PBS (10 min&#160;each).</li>
	<li>Wash the brain 3x&#160;for 20 min in 0.2% Triton X-100 diluted in 0.1 M PBS before dehydrating it in an ascending alcohol series (30%, 50%, 70%, 90%, 95%, 3x 100% ethanol, 20 min each step).</li>
	<li>Embed the dehydrated brain in Methylsalicylate on a microscope slide and seal it with a cover slide.</li>
	<li>Use a confocal laser scanning microscope and scan the brain as optical sections every 2-5 &#181;m using a Harmonic Compound Plan Apochromat objective (10X&#160;0.4 NA immersion). Excite the tissue using 568 nm wavelength for tetramethylrhodamin dextran and a wavelength of 488 nm for the electrode position.</li>
	<li>Reconstruct the stained brain structures and electrode paths from the image stacks in 3D with reconstruction software (e.g. AMIRA or Fiji) (Figure&#160;4).</li>
</ol>

The authors declare that they have no competing financial interests.

This article demonstrates the production and usage of custom designed multi channel micro wire-electrodes. The described electrodes are suitable for recording both single unit and population activity which is especially useful for latency measurements and other temporal response properties of different neurons and different neuropils within a single specimen (for details see1,2,25). Additionally we have shown how to permanently implement the micro&#160;wire electrodes to allow stable long-term recordings in behaving honeybees that last for hours up to days.
Extracellular multi-unit recordings became a favorable tool to achieve high temporal resolution combined with spatial information. In our case, these are either parallel neuronal tracts2 or two different neuropils1. Multiple neurons can be recorded and analyzed at the single neuron level in parallel and at high temporal resolution. Multi-unit recordings were first applied in mammals38 and later also in insects39-41. Substantial progress was achieved with the development and improvement of extracellular multi-channel recording techniques42,43. This, for example, includes the development of new electrodes44 or novel spike sorting and clustering algorithms45. General methods of extracellular multi&#8208;unit recording techniques are well described46-48. The self built electrodes shown in this video additionally can be adapted by adding more microwires per electrode or the micro wires can be twisted to gain measureable constant distances between the tips. Both procedures would, however, lead to decreasing flexibility and increasing thickness of the electrode.
Compared to silicon probes commonly used for extracellular recordings in much larger insects like the hawk moth, locust and cockroach40,49&#8211;51 the described micro&#160;wire-electrodes are smaller, flexible and can cope easily with potential brain movements and, thus, can be reliably used in small social insects like bees and ants that show a much broader behavioral repertoire. Most silicone probes have sharp shank like structures cutting axons and neural tissue along their insertion channel, while the described micro wires are round, flexible and smaller and are hence less harmful to the surrounding tissue which is a clear advantage if the goal is to study long-term plasticity in an intact and behaving animal. Another advantage of micro wire electrodes is their low cost production and the easy handling. Instead of carefully cleaning an expensive silicone probe the electrode wires are freshly cut prior to brain insertion and, therefore, lack problems of congestion. Furthermore it is possible to use more than one micro wire electrode in the same preparation either inserted in different neuropiles1 or tracts2 as we show here. This approach is especially favorable to analyze and compare temporal aspects like response latencies and interactions at different neural processing levels.
We are aware of the fact that an extracellular recorded signal does not reflect single cell activity per se. It is always a compound of voltage activity around the electrode tip. To pinpoint the source of the signal the difference of two neighboring micro&#160;wire channels within one electrode is always computed. Thus the source for the spike signals used to extract single unit activity were always very close to either one or the other electrode channel resulting in easily distinguishable spike waveforms. Signals from farther away, like muscle activity or activity of neighboring neuropils, reach both electrodes at the same time evoking comparable shapes and amplitudes and will be discarded by this procedure. Using the template matching technique of Spike2, we are very confident to obtain single unit activity, which is not the same, but very close to single neuron activity. However, the issue of spike sorting could be avoided by using intracellular recording techniques.
Single cell recordings with either sharp electrodes or patch pipettes allow indepth knowledge about physiological properties of a single neuron. However, due to the small size of insect neurons and their neurites (e.g., less than 1 &#181;m for honeybee PNs 52) only short term recordings are manageable. Furthermore, intra cellular recordings might be invasive and might harm the cell which is possibly another reason for the temporal limitations. In vivo intracellular recordings in insects rarely outlast one hour. A time window which was sufficient for the pioneering work of Martin Hammer53 who recorded intracellular from a single identified neuron, the ventral unpaired maxilar neuron #1 (VUMmx1). He could link its activity directly to the reward pathway. Juliane Mauelshagen54 registered intracellularly the activity of an identified mushroom body extrinsic neuron, the pedunculus extrinsic neuron#1 (PE1) during classical conditioning. The same neuron was in the focus of Menzel and Manz55 when they found LTP after electrical stimulation of Kenyon Cells. However, Okada and colleagues56 could use the intracellularly well characterized spiking pattern (double and triple spikes) for the identification of the PE1 during extracellular recordings. After all a combination of both methods, intracellular recordings from identified neurons and extracellular long term recordings might be a powerful tool for future investigations.
However, using sharp electrodes to record multiple cells (units) simultaneously at different processing levels over many hours up to days to analyze their temporal response relationships and/or even plastic changes is almost impossible.
With the first calcium imaging approaches in the honeybee57,58 using calcium-sensitive dyes the analysis of spatial patterns of odor responses were accessible59-62. However, in many cases the calcium sensitive dyes have to be introduced to the brain tissue via invasive manipulations that again limit the bee&#8217;s life span and intrinsic properties of the analyzed cells. This issue is overcome in other model organisms like the fruitfly using genetically introduced calcium sensors63,64. However, in general, calcium sensors may introduce other limitations as they can act as calcium buffers that likely influence the temporal properties of odor responses. Simultaneous intracellular recordings combined with calcium imaging or computational approaches can prove the proper temporal resolution of imaging processes65,66. However, the temporal resolution of the imaging process itself is rather limited. Optical acquisition systems usually use CCD-imaging with a temporal resolution of 5-20 Hz 67, although 2-Photon-Imaging might be able to acquire faster sequences68. However, increasing sampling rate always goes along with a loss in spatial resolution. Furthermore the calcium-sensitive dyes used in the honeybee undergo bleaching, which also reduces acquisition time69.
Compared to other physiological recording techniques in insects our flexible multi-channel micro-wire electrodes ensure long-time access to single unit and population neuronal activity in behaving bees.
We demonstrated how to use two of these electrodes at different processing stages in the same animal, which facilitates the analysis of temporal coding aspects between the different recording sites. Dependent on the research problem and the model insect the basic method of electrode building demonstrated here is easily extendable and/or can be adapted. For example it is conceivable to use more than the three single wires to produce multi-channel electrodes. In addition, the number of recording sites can be extended and observing temporal aspects of more than two tracts or neuropils is feasible. Our hope is that this method will inspire many scientists and will contribute positively to the understanding of sophisticated neuronal processing in small brains.

Honeybees as well as most other insects heavily rely on olfaction. Amongst others they use olfactory cues for orientation, mating, communication with conspecifics, and foraging. Their well elaborated olfactory system contributes to a rich repertoire of learning behaviors related to floral odor stimuli. These behaviors can be easily studied under controlled laboratory conditions (for review see3-5). Their &#8220;mini brains&#8221; (cp.6) with their relatively small numbers of neurons makes the honeybee a well suited model organism for studying olfactory coding and learning during monitoring of neural activity.
The olfactory system in insects as well as in mammals shows analogous organization to a great extent (for review see7,8). In honeybees about 80,000 receptor neurons9 situated in sensillae along the antennae10,11 translate the environmental odor stimulus into a neuronal signal. Axons from olfactory receptor neurons innervate the antennal lobe (AL), which has a glomerular organization comparable to the vertebrate olfactory bulb. The AL comprises about 164 glomeruli interconnected with each other by about 4,000 local interneurons (LN) (for review see12). Especially in the honeybee it has been shown recently that LNs provide patchy lateral connectivity and that different subpopulations possess elemental and configural olfactory coding properties13,14 . The AL was shown to be subdivided into a ventral and a dorsal hemi lobe giving rise to the medial and the lateral antennal lobe tract (m- and l-ALT; formerly termed m- and l-APT for medial- and lateral antennal lobe protocerebral tract15-17). Here a new tract terminology introduced by a recent effort for a unified nomenclature of the insect brain will be used18. Both ALTs (l- and m-ALT) combine either 410 (l-ALT) or 510 (m-ALT) uniglomerular projection neurons (PN), respectively15,16,19. PNs of both tracts have recently been shown to code odors in parallel2&#160;(for review see17,20), and both tracts synaptically form divergent connections with Kenyon Cells (KC), the mushroom body (MB) principal neurons. Each MB contains about 172,000 KCs21-23. The MBs are known to be involved in stimulus integration, learning, and memory formation. The axo dendrites of KCs form the peduncle (the mushroom&#180;s stem), which has two main output regions: the vertica &#160;or alpha- lobe and the horizontal &#160;or beta-lobe22,24. The output of the MB converges to only about 400 extrinsic neurons (EN)24. ENs responsible for olfactory information processing mostly innervate the ventral aspect of the vertical lobe22. Recently, it has been shown that ENs recorded at this area encode the odor reward association25.
Temporal aspects within the olfactory system of insects as well as vertebrates have become an important and significant aspect as a potential coding principle26-29. To be able to simultaneously record multiple neurons from different sites at high temporal resolution, we established double multi unit recording techniques using customized multi channel wire electrodes introduced to different target regions in the bee&#8217;s olfactory system. This approach enables us to analyze and compare temporal processing in the honeybee olfactory system at the level of single neurons and populations of neurons either between parallel olfactory pathways, the dual olfactory pathway2 or between different subsequent neuropils1. Recently with a similar experimental approach in the locust olfactory system30 using a different configuration of electrodes were able to analyze spatiotemporal coding mechanism for background-invariant odor recognition31. Thus, the established dual recordings enable gathering spatial information about simultaneous neuronal activity profiles.
Compared to the broader spatial sampling obtained from calcium imaging this method allows recording from two spots only. However, the advantage compared to calcium imaging techniques is the high temporal precision of action potential recordings, which cannot be provided by either conventional CCD imaging or 2-photon&#160;imaging acquisition. The extracellular electrodes described here are permanently implanted and fixed relative to the brain and head capsule avoiding electrode drift. This is a clear advantage compared to the usage of sharp intracellular electrodes. Another advantage compared to intracellular recordings and calcium imaging is the extended neural observation time ranging from many hours up to days. This is an important prerequisite to investigate neural correlates of learning and memory formation. Additional benefits of multi unit recordings are further outlined in the discussion section.
In this methodological overview the manufacturing procedure of custom design wire electrodes will be shown, adapted from32,33 and suited for long term multi unit recordings in the honeybee brain. Additionally, an example how these types of electrodes are permanently implanted at two different recording sites within the honeybee olfactory system to record simultaneously the l- and the m-ALT over long periods of time to allow many stimulation protocols is shown2. For verification of the recording positions an example and protocol for staining and post recording visualization of the recording sites is provided.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Paraffin oil</td>
			<td>Fluka</td>
			<td>76235</td>
			<td></td>
		</tr>
		<tr>
			<td>Odors</td>
			<td>Sigma Aldrich&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td></td>
			<td></td>
			<td>pH 7.2</td>
		</tr>
		<tr>
			<td>4% Formaldehyde</td>
			<td>ThermoScientific</td>
			<td>28908</td>
			<td>Methanol free</td>
		</tr>
		<tr>
			<td>Triton X</td>
			<td>BioChemica</td>
			<td>A1388</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylsalicylate</td>
			<td>Roth</td>
			<td>4529.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Tetramethylrhodamin dextran, 10,000 MW (Microruby)</td>
			<td>Invitrogen</td>
			<td>D7162</td>
			<td>keep dark</td>
		</tr>
		<tr>
			<td>Alexa 488 hydrazide</td>
			<td>Invitrogen</td>
			<td>A-10436</td>
			<td>keep dark</td>
		</tr>
		<tr>
			<td>Alexa 568 hydrazide</td>
			<td>Invitrogen</td>
			<td>A-10437</td>
			<td>keep dark</td>
		</tr>
		<tr>
			<td>Bee Ringer Solution</td>
			<td>see 2</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Polyurethane-coated copper wire</td>
			<td>Elektrisola</td>
			<td>15&#181;m diameter &#38; P155 insulation</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental Wax&#160;</td>
			<td>Densply Detrey</td>
			<td>64103015S1</td>
			<td>moderate melting point</td>
		</tr>
		<tr>
			<td>Dental Wax</td>
			<td>Flexaponal&#160;</td>
			<td>124-202-00</td>
			<td>low-melting Wax</td>
		</tr>
		<tr>
			<td>KWIK SIl</td>
			<td>WPI</td>
			<td>03L</td>
			<td></td>
		</tr>
		<tr>
			<td>18 Pin Socket</td>
			<td>Conrad Electronic</td>
			<td>189634-62</td>
			<td></td>
		</tr>
		<tr>
			<td>Hot melting glue</td>
			<td>Conrad Electronic</td>
			<td>827673</td>
			<td></td>
		</tr>
		<tr>
			<td>soldering needle</td>
			<td>Conrad Electronics</td>
			<td>830283</td>
			<td>12 V</td>
		</tr>
		<tr>
			<td>Soldering terminal lug&#160;</td>
			<td>Conrad Electronic</td>
			<td>531901</td>
			<td></td>
		</tr>
		<tr>
			<td>Glaselectrodes</td>
			<td>WPI</td>
			<td>1B100F-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Minutien Pins</td>
			<td>Fine Science Tools</td>
			<td>26002-20</td>
			<td>V2A 0.2 x 12 mm</td>
		</tr>
		<tr>
			<td>switchable headstage</td>
			<td>Tucer Davis Technologies</td>
			<td>SH16</td>
			<td></td>
		</tr>
		<tr>
			<td>Headstage connection module</td>
			<td>NPI</td>
			<td>INT-03M</td>
			<td></td>
		</tr>
		<tr>
			<td>Amplifier Module</td>
			<td>NPI</td>
			<td>PDA-2F</td>
			<td></td>
		</tr>
		<tr>
			<td>Data Acquisition boards</td>
			<td>National Instruments</td>
			<td>NI-6123, Ni-6143</td>
			<td></td>
		</tr>
		<tr>
			<td>Acquisition Software</td>
			<td>National Instruments</td>
			<td>Lab View 8.2</td>
			<td>custom design</td>
		</tr>
		<tr>
			<td>Spike-Sorting&#160;</td>
			<td>CED&#160;</td>
			<td>Spike 2 v7.11</td>
			<td></td>
		</tr>
		<tr>
			<td>Matlab</td>
			<td>Mathworks</td>
			<td>R2008B</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Leitz</td>
			<td></td>
			<td>manual</td>
		</tr>
		<tr>
			<td>AG-wires</td>
			<td>WPI</td>
			<td>AGT05100</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal laser scanning microscope</td>
			<td>Leica</td>
			<td>TCS SP2 AOBS</td>
			<td></td>
		</tr>
		<tr>
			<td>AMIRA</td>
			<td>Mercury Computer Systems&#160;</td>
			<td>2/5/2000</td>
			<td></td>
		</tr>
	</tbody>
</table>

simultaneous long-term recordings at two neuronal processing stages in behaving honeybees

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent and divergent anatomical connections including feed forward and feedback wiring. Furthermore, information of the same origin is partially sent via parallel pathways to different and sometimes into the same brain areas. To understand the evolutionary benefits as well as the computational advantages of these wiring strategies and especially their temporal dependencies on each other, it is necessary to have simultaneous access to single neurons of different tracts or neuropiles in the same preparation at high temporal resolution. Here we concentrate on honeybees by demonstrating a unique extracellular long term access to record multi unit activity at two subsequent neuropiles1, the antennal lobe (AL), the first olfactory processing stage and the mushroom body (MB), a higher order integration center involved in learning and memory formation, or two parallel neuronal tracts2 connecting the AL with the MB.&#160;The latter was chosen as an example and will be described in full. In the supporting video the construction and permanent insertion of flexible multi&#160;channel wire electrodes is demonstrated. Pairwise differential amplification of the micro&#160;wire electrode channels drastically reduces the noise and verifies that the source of the signal is closely related to the position of the electrode tip. The mechanical flexibility of the used wire electrodes allows stable invasive long&#160;term recordings over many hours up to days, which is a clear advantage compared to conventional extra&#160;and intracellular in vivo recording techniques.

&#34;The present protocol allows simultaneous recordings at two different processing stages within individual honeybees and additionally allows to test underlying mechanisms of learning and memory via e.g., PER conditioning within restrained honeybees.&#34;&#160;This is a prerequisite for analyzing temporal aspects of neuronal processing. The method is easily adaptable for different scientific approaches to unravel the neuronal network of the bee&#8217;s olfactory system. For example this method is used (i) to analyze temporal processing of PNs within the dual olfactory pathway of the honeybee, the l- and m-ALT PNs (Figure&#160;5). In Figure 5A one example of an l-ALT PN simultaneously recorded with a PN of the m-ALT is given as ten trial average and illustrates their response strength and latency in respect to five different odor concentrations as color coded heat plot. In an average of seven bees with 11 l- and 13 m-ALT PNs (Figures 5B, 5C) illustrates that both the response strength as well as the response latency, to a certain extent, reflects odorant concentration. Thereby PNs in this example increase their response strength while with increasing odorant concentration their response latency declined (Figures 5B, 5C). This result is rather limited and only valid for the analyzed odorant, but is still consistent with recent computational models of the AL37. Whether odor concentration coding in the bee&#8217;s AL underlies non linear computations or underlies other coding properties still needs to be analyzed in future. Furthermore the method can be used (ii) to compare temporal aspects in the population activity at two subsequent processing stages, the AL- and MB-output (Figure&#160;6). Principal component analysis (PCA) illustrates that odor computation is prolonged and outlast the entire odor presentation at the PN level whereas in ENs only the odor on&#160;and off set were represented in the population activity (Figure&#160;6). Thereby the EN population reached their maximal activity already at a time point when the PN activity is still developing (cp. Movie 1).
<img alt="Figure 1" fo:content-width="4in" src="/files/ftp_upload/51750/51750fig1highres.jpg" width="400" /> 
Figure 1.&#160;Manufacturing three-channel micro-wire electrodes.&#160;A1) The endings of three wires are soldered to an IC pin connector in Positions 11, 13 and 16. A2) Four soldering lugs are screwed onto a Plexiglas plastic base plate; a minutien pin is inserted into the tip of a glass capillary which is then attached to the plastic plate. A3) The plastic plate is glued on top of an IC pin connector and the free end of each wire is soldered to one of the top soldering lugs. B1) To equip the electrode holder with the fine copper wires, the capillary has to be taken out once more. B2) The base of the holder is then fixed in a custom made aligning device. Three copper micro wires are aligned along the groove and fixed with adhesive tape at each end. B3) The parallel micro wires are glued together with dental wax and the capillary is put back in place (long arrows); the glued micro wires are then attached to the capillary with dental wax and its ends are cut off (short arrows). B4) The three loose ends of the copper wires are soldered to the three top soldering lugs thus bringing them into electrical contact with the IC pin connector. C1) Two fully assembled electrodes can be connected to each other for use with a single headstage. C2) For this purpose the pins of one electrode (left, slave) are connected via insulated wires to the other electrode (right, master), which is connected to the head stage; the headstage connected electrode can also collect input from a muscle (M17) and reference electrode (Ref).
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51750/51750fig2highres.jpg" width="500" /> 
Figure 2.&#160;Preparation and permanent electrode insertion into the bee brain.&#160;A) A bee is inserted into a Plexiglas holder after immobilization on ice. Antennae with Flagellum (FL) and Scapus (SC) are indicated. B) The head and antennae are fixed using dental wax. C) The head capsule is shaved and the bee is fed with sugar water. D) The head capsule is opened. E) After removing glands and trachea from the top of the brain, the different neuropiles and major landmarks can easily be distinguished. The trajectories of the ALTs are indicated together with a mark where the electrodes are inserted. (MB: Mushroom body, AL: Antennal lobe, OL: Optical lobe, &#945;: Alpha lobe or vertical lobe, ALT: antennal lobe tract, E1: electrode insertion side for m-ALT recordings, E2: electrode insertion side for l-ALT recordings). F) Reference (Ref) and muscle electrodes (M17) are inserted into the head capsule through little holes in the cuticle or the compound eye. G) The wire electrodes are inserted into the brain at the appropriate sites. H) After fixing the electrodes in place using two component silicon, the bee still shows PER and can be conditioned (e.g., using sugar water).
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51750/51750fig3highres.jpg" width="500" /> 
Figure 3.&#160;Extracellular recording at two neural tracts and single unit extraction (spike sorting). A) Schematic drawing of the experimental setup. The bee is fixated in a plexiglas holder. Odor stimulation is provided via a glass tube. Two electrode shanks are recording from the exposed bee brain. B) Simultaneous recordings from the l- and m-ALT PNs (green and purple traces) showing excitatory responses on both tracts to a 500 msec odor stimulation of honey in water solution at a concentration of 1:100 at 33 &#176;C. Each plotted line represents the differentiated channels as color-coded label from the electrode soldering lugs in A. Bar: 50 &#181;V. C) After spike sorting procedures single action potentials are sorted and color coded. Overlay of the sorted units illustrates the separation of the waveforms. D) Spike interval histogram indicates the separation quality of the sorted units as proof of adequate spike sorting. Note there is no spike within the unit&#8217;s refractory period. E) Two views (E1, E2) from a 3D clustering of the sorted unit with principal component analysis which indicates the distance of the sorted units to each other. Circles indicate the 2.5 fold Mahalanobis distance which resembles the SD in space and indicates a significant differentiation of the clusters in the principal component space F) Color coded units depict the action potentials visible in a magnification of three channels from one tract recording. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51750/51750fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51750/51750fig4highres.jpg" width="500" /> 
Figure 4.&#160;Post-recording visualization and 3D-reconstruction of the recording position.&#160;A) Projection view of ortho-slices along the z-axes with maximum intensity alignment of an antero&#160;and retrograde backfilling of the uniglomerular projection neurons projecting from the AL to the MB and LH. The intracellular tracer Microruby (tetramethylrhodamin dextran) was inserted into the AL after the recording experiments. Stained glomeruli inside the AL proof proper PN staining. B) Projection view of ortho slices along the z -axes with maximum intensity alignment from a staining of the two electrodes with Alexa hydrazide 488 indicating the electrode placement for the m-ALT (E1, arrow) and l-ALT (E2, arrow). The tracer Alexa Hydrazide 488 migrates to the electrode surrounding tissue and stains the electrode insertion site. Note, the prominent staining in the AL is a superficial artefact. C) 3D Reconstructions of the stained target cells (PNs) and the electrode insertion site from A,B (right side) together with a schematic overview of the honeybee olfactory system (left side) with the indication of l- and m-ALT trajectories. Note, only uniglomerular PN tracts are shown. AN: antennal nerve, AL: antennal lobe, LH: lateral horn, MB: mushroom body, E1,E2: electrode insertion sites, m-ALT: medial antennal lobe tract, l-ALT: lateral antennal lobe tract, c: caudal, r: rostral, m: medial, l: lateral. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51750/51750fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="3in" src="/files/ftp_upload/51750/51750fig5highres.jpg" width="300" /> 
Figure 5.&#160;Example of odor concentration coding within the dual olfactory pathway. A) Heat plots illustrate the response of a single l-ALT (green) and single m-ALT PN (purple) acquired simultaneously from an individual honeybee in response to the odorant hexanal at increasing odor concentrations (from 1: 10-6 to 1:100). Each line is an average of ten trial stimulation. The firing rate is shown as relative intensity change which is the firing rate in relation to the subtracted spontaneous activity. B) Population response latency of 11 l- and 13 m-ALT PNs from 7 recorded bees. In each bee the PNs from both tracts were recorded simultaneously. The population response latency shows a decreasing latency with increasing odor concentration in PNs from both tracts. The odor response onset at the antennae at 99 msec was recorded via electroantennograms and is subtracted from the PN response latencies. Note that at the lowest concentrations responses are too weak for latency measurements and, therefore, were excluded. C) Population response firing rate from same PNs as in B. With increasing odor concentration the response strength is increasing. The l-ALT shows a stronger response strength. In B,C the mean and SD are given.
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/51750/51750fig6highres.jpg" width="500" /> 
Figure 6.&#160;Comparing population activity at two subsequent processing stages along the honeybee&#8217;s olfactory pathway.&#160;Data were recorded in 20 animals which were stimulated with 1-hexanol and 2-octanone. A) Each line represents the false color coded mean firing rate of one projection neuron (PN) calculated across 10 odor repetitions of 1-hexanol. Odor presentation starts at time 0 and lasted three seconds. B) Shows the same as in A) but for mushroom body extrinsic neurons (EN). The matrix shown in A) can be seen as a PN population vector during odor stimulation with 1-hexanol. We calculated the same kind of population vector during odor stimulation with 2-octanone and used both vectors in a principal component analysis (PCA) keeping the temporal dimension. C)&#160;The first three principal components (PC1, 2 and 3) were plotted against each other to illustrate the odor separation in the PN ensemble activity at the antennal lobe output. The time before odor onset is marked in black. Activity during three seconds of stimulation with 1-hexanol is shown in blue. The activity during stimulation with 2-octanol is shown in red. Furthermore, we show 1.5 seconds (post) of the activity after odor of set of 1-hexanol (light-blue) and 2-octanonen (pink). Note that at the PN ensemble level, both odors evoking very distinct trajectories settle in a &#8220;fixed point&#8221; which outlasts the whole odor stimulation period. Only after odor offset the trajectories move back to the baseline activity without odor stimulation. D) The same analysis was done at the EN ensemble level representing the activity at the mushroom body output. Compared to the PN activity odors evoke a less distinct trajectory. Furthermore a &#8220;fixed point&#8221; is not observable. The initially odor induced trajectories intermingle with baseline activity although the odor is still present. Only the odor offset evoked an additional trajectory.
<a href="/files/ftp_upload/51750/Movie_R3_Brill_et_al._JoVE.avi" target="_blank">Movie 1.</a> Time resolved evaluation of an odor induced trajectory after principal component analysis of a PN-population vector (left) and an EN population vector (right; cp. Figure&#160;6). The upper parts include the first three principal components (PC1, 2 and 3) plotted against each other. The lower panels illustrate the evaluation of PC1, 2 and 3 over time. Odor stimulation is marked by the gray bar. All panels were synchronized. Note that the EN population activity starts slightly before the PN population activity,&#160;a phenomenon which seems to be contraintuitive but can be explained by the connectivity and properties of the involved layers, which is discussed earlier1.

Watch this Scientific Journal Video about Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees at JoVE.com

Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees

Simultaneous extracellular long term recordings from two different brain neuropiles or two different anatomical tracts were established in honeybees. These recordings allow the investigation of temporal aspects of neuronal processing across different brain areas at the single neuron as well as at the ensemble level in a behaving animal.

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent ...

simultaneous-long-term-recordings-at-two-neuronal-processing-stages

Nanyang Technological University

Research

JoVE Journal

Neuroscience

12.8K Views.  University of Würzburg. Simultaneous extracellular long term recordings from two different brain neuropiles or two different anatomical tracts were established in honeybees. These recordings allow the investigation of temporal aspects of neuronal processing across different brain areas at the single neuron as well as at the ensemble level in a behaving animal.

Video: Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees

Automated Sholl Analysis of Digitized Neuronal Morphology at Multiple Scales

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3. Specifically, it affects the ability of neurons to receive inputs from other cells2 and contributes to the propagation of action potentials4,5. The morphology of the neurites also affects how information is processed. The diversity of dendrite morphologies facilitate local and long range signaling and allow individual neurons or groups of neurons to carry out specialized functions within the neuronal network6,7. Alterations in dendrite morphology, including fragmentation of dendrites and changes in branching patterns, have been observed in a number of disease states, including Alzheimer's disease8, schizophrenia9,10, and mental retardation11. The ability to both understand the factors that shape dendrite morphologies and to identify changes in dendrite morphologies is essential in the understanding of nervous system function and dysfunction.
Neurite morphology is often analyzed by Sholl analysis and by counting the number of neurites and the number of branch tips. This analysis is generally applied to dendrites, but it can also be applied to axons. Performing this analysis by hand is both time consuming and inevitably introduces variability due to experimenter bias and inconsistency. The Bonfire program is a semi-automated approach to the analysis of dendrite and axon morphology that builds upon available open-source morphological analysis tools. Our program enables the detection of local changes in dendrite and axon branching behaviors by performing Sholl analysis on subregions of the neuritic arbor. For example, Sholl analysis is performed on both the neuron as a whole as well as on each subset of processes (primary, secondary, terminal, root, etc.) Dendrite and axon patterning is influenced by a number of intracellular and extracellular factors, many acting locally. Thus, the resulting arbor morphology is a result of specific processes acting on specific neurites, making it necessary to perform morphological analysis on a smaller scale in order to observe these local variations12.
The Bonfire program requires the use of two open-source analysis tools, the NeuronJ plugin to ImageJ and NeuronStudio. Neurons are traced in ImageJ, and NeuronStudio is used to define the connectivity between neurites. Bonfire contains a number of custom scripts written in MATLAB (MathWorks) that are used to convert the data into the appropriate format for further analysis, check for user errors, and ultimately perform Sholl analysis. Finally, data are exported into Excel for statistical analysis. A flow chart of the Bonfire program is shown in Figure 1.

We have developed a computer program to analyze neuronal morphology. In combination with two existing open source analysis tools, our program performs Sholl analysis and determines the number of neurites, branch points, and neurite tips. The analyses are performed so that local changes in neurite morphology can be observed.

Neuronal morphology plays a significant role in determining how neurons function and communicate1-3.  Specifically, it affects the ability of neurons to ...

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

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

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

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

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

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

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

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

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

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

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

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

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

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

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

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

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

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

Generation and Long-term Maintenance of Nerve-free Hydra

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The interstitial cells can be eliminated through two consecutive treatments with colchicine, a plant-derived toxin that kills dividing cells, thus erasing the potential for renewal of the differentiated cells that are derived from the interstitial stem cells. This allows for the generation of Hydra that lack nerve cells. A nerve-free polyp cannot open its mouth to feed, egest, or regulate osmotic pressure. Such animals, however, can survive and be cultured indefinitely in the laboratory if regularly force-fed and burped. The lack of nerve cells allows for studies of the role of the nervous system in regulating animal behavior and regeneration. Previously published protocols for nerve-free Hydra maintenance involve outdated techniques such as mouth-pipetting with hand-pulled micropipette tips to feed and clean the Hydra. Here, an improved protocol for maintenance of nerve-free Hydra is introduced. Fine-tipped forceps are used to force open the mouth and insert freshly killed Artemia. Following force-feeding, the body cavity of the animal is flushed with fresh medium using a syringe and hypodermic needle to remove undigested material, referred to here as "burping". This new method of force-feeding and burping nerve-free Hydra through the use of forceps and syringes eliminates the need for mouth-pipetting using hand-pulled micropipette tips. It thus makes the process safer and significantly more time efficient. To ensure that the nerve cells in the hypostome have been eliminated, immunohistochemistry using anti-tyrosine-tubulin is conducted.

Generation and Long-term Maintenance of Nerve-free Hydra

Through a double treatment with colchicine, a plant-derived toxin that kills dividing cells, nerve-free Hydra vulgaris can be generated. These Hydra cannot feed or egest on their own. This paper describes an improved method for long-term maintenance of nerve free Hydra vulgaris in the laboratory.

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

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 ...

Long-term Sensory Conflict in Freely Behaving Mice

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning in mice. By permanently wearing a device fixed on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages. Therefore, this protocol readily enables the study of the visual system and multisensory interactions over an extended timeframe that would not be accessible otherwise. In addition to lowering the experimental costs of long-term sensory learning in naturally behaving mice, this approach accommodates the combination of in vivo and in vitro experiments. In the reported example, video-oculography is performed to quantify the vestibulo-ocular reflex (VOR) and optokinetic reflex (OKR) before and after learning. Mice exposed to this long-term sensory conflict between visual and vestibular inputs presented a strong VOR gain decrease but exhibited few OKR changes. Detailed steps of device assembly, animal care, and reflex measurements are hereby reported.

The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning. By permanently wearing a fixed device on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages.

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for ...