We would like to thank Profs. Huibert Mansvelder and Bert Sakmann for extensive support, Dr. Marcel Oberlaender for fruitful discussions and providing neuronal tracing, and Brendan Lodder for technical assistance. Data was acquired using the ntrode VI for LabView, generously provided by R. Bruno (Columbia Univ., NY, USA). This research was supported by the Max Planck Society and the Bernstein Center for Computational Neuroscience, Tuebingen (funded by the German Federal Ministry of Education and Research (BMBF; FKZ: 01GQ1002))( R.T.N.), Center for Neurogenomics and Cognitive Research (CNCR), Neuroscience Campus Amsterdam (NCA), funding to C.P.J.d.K. (NWO-ALW #822.02.013 and ENC-Network #p3-c3) and the VU University Amsterdam.

1. Preparation of the Animal
All experimental procedures are carried out in accordance with the Dutch law and after evaluation by a local ethical committee at the VU University Amsterdam, The Netherlands.
<ol>
	<li>Anesthetize a Wistar rat (P25-P45, &#9794;/&#9792;) with isoflurane (2-3% in oxygen) and subsequently with urethane (20% in 0.9% NaCl, 1.6-1.7 g/kg) by intraperitoneal injection. Assess the depth of the anesthesia by monitoring pinch withdrawal, eyelid reflexes, and vibrissae movements.</li>
	<li>Administer an additional dose of urethane (10% of initial dose) in case the animal does not reach sufficient anesthetic depth or whenever vibrissae twitches are observed during the course of the experiment.</li>
	<li>Place the anesthetized animal on the stereotaxic frame equipped with a heating pad, insert rectal temperature probe and maintain the animal&#8217;s body temperature at 37.5&#177;0.5 &#176;C by a heating pad.</li>
	<li>Position the animal&#8217;s head in the stereotaxic frame; trim the hair on top of the head using scissors to render the skin visible.</li>
	<li>Inject 100 &#181;l 1% lidocaine (in 0.9% NaCl) subcutaneously for local anesthesia at the operation site and wait 3-5 min. Remove the skin covering the surface of the skull by cutting in the caudal-to-rostral direction and clean the remaining tissue.</li>
	<li>Clean the skull extensively with 0.9% NaCl and absorbent swabs.</li>
	<li>Determine the coordinates of the target area on the left hemisphere (for primary somatosensory cortex (S1): 2.5 mm posterior and 5.5 mm lateral to bregma). Mark the location on the skull with a surgical skin marker pen.</li>
	<li>Scrape off the bone gently by a dental drill from the region of interest until the bone becomes transparent and blood vessels are clearly visible.</li>
	<li>Make a 0.5 mm x 0.5 mm craniotomy by cutting a window in the thinned skull with a scalpel (#11), avoiding damage to the dura mater and blood vessels. Optional: mark the edges of the craniotomy using a surgical skin marker pen to improve visibility.</li>
	<li>Apply a thin layer of superglue on the dry skull and then dental cement to construct a bath (i.e. recording chamber) enclosing the craniotomy.</li>
	<li>Trim the whiskers at the contralateral side to exactly 5 mm from the whisker follicle. Optional: highlight the trimmed whiskers with black mascara.</li>
</ol>
2. Juxtasomal Recordings and Biocytin Labeling
<ol>
	<li>Make patch pipettes of borosilicate glass with inside tip diameter of ~1 &#956;m to obtain electrodes with 3-5 M&#937; resistance. Note: The ideal pipette morphology for the juxtasomal recording is a gradual slender taper, a low cone angle, and a ~1 &#956;m tip (Figure 1).</li>
	<li>Depending on the recording depth (with respect to the pial surface), the taper dimensions of the recording electrode should be adjusted. Critically important is that the taper diameter should be less than 75 &#956;m at point of entry in the brain to avoid mechanical damage or stress to the recording area. This indicates that for superficial cortical recordings, a taper of 300-500 &#956;m with an outer diameter of maximally 75 &#956;m is required (Figure 1A) whereas recordings from relatively deep cortical layers require a longer taper of 1,500-1,800 &#956;m, again with maximal outer diameter of 75 &#956;m (at 1,800 &#956;m from the electrode tip).</li>
	<li>Load the patch pipette with normal rat ringer (NRR) supplemented with 2% biocytin (see Table 1 for solutions and&#160;reagents) and mount the patch pipette on a head stage fixed to a micromanipulator.</li>
	<li>Set the angle of the electrode holder to 34&#176; with respect to the sagittal plane to specifically target the D2 column of rat S1.</li>
	<li>Connect the head stage to an amplifier in bridge-mode (i.e. current clamp).</li>
	<li>Position the patch pipette in close proximity to the craniotomy. Fill the bath with 0.9% NaCl and determine the electrode resistance which should be between 3-5 M&#937;, assessed by applying a square pulse with positive current injection (1 nA, 200 msec on/off).</li>
	<li>Establish overpressure of 100-150 mbar and advance the patch pipette in 1 &#181;m steps while applying positive current as square pulses (1 nA, 200 msec on/off). Monitor the robust resistance change upon establishing contact with the dura mater. At this point, set the coordinates of the micromanipulator to &#8216;zero&#8217; to allow accurate depth measurements.</li>
	<li>Advance in 1 &#181;m step-mode until the patch pipette penetrates the dura mater indicated by a sudden drop in electrode resistance. Remove the holding pressure of the pipette.</li>
	<li>Search for single units while advancing in 1 &#181;m steps. Monitor the electrode resistance continuously by applying 200 msec on/off pulses. An increase in electrode resistance typically indicates the proximity of a single neuron. Advance the electrode until positive action potential (AP) waveforms of ~2 mV are recorded (Figures 2A and 2B).</li>
	<li>Optional: Record the spontaneous spiking pattern of the neuron and determine whisker-evoked spiking by, for instance, caudally deflecting individual whiskers for 200 msec at an angle of 3.3&#176; using a piezoelectric device18.</li>
	<li>Advance the electrode until the resistance is 25-35 M&#937; and spikes have amplitudes of 3-8 mV to obtain optimal conditions of juxtasomal filling. Start the juxtasomal filling by applying square pulses of positive current (1 nA, 200 msec on/off). Slowly and gradually increase the current by steps of 0.1 nA while monitoring the AP waveform and frequency (Figures 2A and 2B).</li>
	<li>Monitor the membrane opening as a clear increase in AP frequency during the on-phase of the block pulse (Figure 2C). The spike waveform during filling shows an increased width and reduced after-hyperpolarization (Figures 2C and 2D). Additional parameters include increased noise or a small (1-5 mV) negative DC shift.</li>
	<li>Increase or decrease the current (1 nA) while filling to maintain stable biocytin infusion. Reduce or stop the current pulses upon sudden increase of the AP frequency to avoid toxicity by excess influx of extracellular ions, such as sodium ions. Note: every neuron will respond differently to the current injection and juxtasomal labeling parameters need to be adjusted depending on individual recording conditions.</li>
	<li>Closely monitor the signal after stopping the current injection. The spike waveform after a filling session is usually broadened and shows a strongly reduced after hyperpolarization. Wait for recovery of the neuron, which is apparent when the spike waveform returns to its original properties (i.e. presence of normal after-hyperpolarization, Figure 2).</li>
	<li>Repeat biocytin filling sessions after complete recovery of the neuron to increase biocytin load for improved staining quality.</li>
	<li>Retract the patch pipette in steps of 1 &#181;m until the spike amplitude decreases to reduce any mechanical stress to the cell. Wait at least 1 hr for the biocytin to diffuse intracellularly while closely monitoring the animal&#8217;s body temperature and breathing.</li>
</ol>
3. Perfusing the Animal and Removing the Brain
<ol>
	<li>Prepare the perfusion setup, rinse, and preload the tubing with 0.9% NaCl.</li>
	<li>Position the animal on a surgical tray and secure with standard labeling tape. Ensure sufficient depth of the anesthesia; foot pinch and eyelid reflexes should be absent.</li>
	<li>Make a medial to lateral incision (5-6 cm) through the abdominal wall just beneath the rib cage and proceed in posterior-anterior direction to expose the sternum. Pull the sternum anteriorly and carefully separate the liver from the diaphragm.</li>
	<li>Make a small incision in the diaphragm, cut through the lower ribs and continue the incision along the entire length of the abdominal cavity to expose the heart.</li>
	<li>Remove the pericardium.</li>
	<li>Insert the needle into the left ventricle and make an incision in the right atrium. Perfuse with 0.9% NaCl (~8 ml/min) until decoloration of the liver is complete.</li>
	<li>Switch the infusion to 4% paraformaldehyde (PFA) until stiffness of front paw and lower jaw is apparent.</li>
	<li>Decapitate the rat using a pair of scissors</li>
	<li>Remove the remaining neck muscles and expose the skull completely.</li>
	<li>Position the scissors in the brain stem on the dorsal side and cut the bone carefully along the sagittal suture, maintaining the dorsal position.</li>
	<li>Remove the bones from both sides of the sagittal suture to expose the brain using forceps. Carefully remove the dura to avoid damage.</li>
	<li>Carefully insert a blunt spatula to the ventral side of the brain and remove the brain gently.</li>
	<li>Post-fix the entire brain overnight in PFA at 4 &#176;C. Switch the brain to 0.05 M phosphate buffer (PB) and store at 4 &#176;C.</li>
</ol>
4. Slicing the Brain in Tangential Sections
<ol>
	<li>Take the brain out of the 0.05 M PB and put it on a filter paper facing anterior. Use a sharp razor blade to cut off the cerebellum along the coronal plane and separate the hemispheres by cutting along the mid sagittal plane.</li>
	<li>Apply superglue on the mounting platform and mount the left hemisphere on its sagittal plane with anterior facing right. Secure the mounting platform at an angle of 45&#176; on a vibratome and submerge the brain in 0.05 M PB.</li>
	<li>Secure a razor blade on the vibratome and make sure that the first contact with the brain surface is in the middle of the anterior-posterior plane of the hemisphere. Cut 24 100 &#181;m sections and collect them in a 24-well plate containing 0.05 M PB.</li>
</ol>
5. Histological Procedures
<ol>
	<li>Perform histological protocols for the cytochrome oxidase staining24 and the avidin-biotin-peroxidase method according to previously described methods25. Optional: visualize biocytin using fluorescent avidin/streptavidin-Alexa conjugates. This additionally allows double-staining with retrograde or anterograde tracing techniques.</li>
	<li>Wash sections 5 x 5 min with 0.05 M PB and prepare the 3,-3&#8217;-diaminobenzidine tetrahydrochloride (DAB)-containing solution for the cytochrome oxidase staining to visualize barrels in layer 4 (L4) of S1 (see Table 2 for solutions and reagents). Incubate sections 6-12 from the pia in the preheated solution for 30-45 min at 37 &#176;C.</li>
	<li>Rinse sections with 0.05 M PB for 6 x 5 min and quench endogenous peroxidase activity by incubating all sections in 3% H2O2 in 0.05 M PB for 20 min at room temperature (RT).</li>
	<li>Rinse sections with 0.05 M PB for 5 x 10 min. Incubate sections in ABC-solution (see Table 3 for solutions and reagents) overnight at 4 &#176;C.</li>
	<li>Rinse sections with 0.05 M PB for 5 x 10 min and prepare the DAB-solution to visualize the biocytin-filled neuron (see Table 4 for solutions and reagents). Incubate sections in filtered solution for 45-60 min at RT.</li>
	<li>Rinse sections with 0.05 M PB for 5 x 10 min. Mount sections on microscope slides and cover slip with Mowiol (see Table 5 for solutions and reagents).</li>
	<li>Determine labeling quality using light microscopy.</li>
</ol>

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The authros have&#160;nothing to disclose.

The juxtasomal method allows recording in vivo action potential spiking from single units across behavioral conditions (anesthetized, awake head-fixed or freely moving) with the option of biocytin-labeling the recorded neuron for post&#160;hoc cell type classification and/or 3D reconstruction. The major advantage is to obtain physiological parameters in the extracellular (thus noninvasive) configuration, yet being able to label the neuron intracellularly with biocytin16,17,32. In addition to ...

Neuronal networks consist of multiple cell types, characterized by highly specific morphological and physiological properties1-7. As a consequence, individual cell types perform specialized tasks within the network (see for instance&#160;Gentet&#160;et al.8 and&#160;Burgalossi&#160;et al.9). We are only beginning to understand cell type-specific functions across neuronal networks and much is still to be discovered. To this end, many labs are implementing experimental approaches that allow the analysis of morphological properties of the same neuronal population from which physiological parameters have been obtained1,10-15. Here, we demonstrate the juxtasomal labeling technique16,17 which involves electrophysiological recordings using conventional patch pipettes in the extracellular (thus noninvasive) configuration in combination with electroporation of the recorded neuron with biocytin. The major advantage of this approach is that the noninvasive nature ensures that action potential spiking of individual neurons is recorded without altering (e.g. dialyzing) the intracellular content of the cell. Followed by electroporation, the juxtasomal approach provides the option of post&#160;hoc cell identification and reconstruction to link function (physiology) to structure (morphology). Typically, morphological reconstruction involves reconstruction of dendritic and axonal morphology which can be extended to quantification of spine and/or bouton densities or even reconstruction of neuronal morphology at nanometer resolution using electron microscopy. The juxtasomal recording technique can be used for in vivo recordings of various cell-types across cortical layers or in sub-cortical areas in a range of species, although most studies have applied the technique in small rodents such as mice or rats. Our research is focused on recording and labeling neurons from rat primary somatosensory cortex (S1) and involves visual identification of recorded neurons18, dendritic reconstructions in combination with precise registration in a standardized reference frame to reverse engineer cortical networks4,19 and detailed reconstruction of axonal architecture to characterize cell type-specific local and long-range projection targets20.
Compared to alternative in vivo recording techniques (intracellular or whole-cell), juxtasomal recordings are relatively stable and can therefore be applied across behavioral states including anesthetized21,22, sedated14, awake head-fixed23, or even freely-moving animals9. Here, we show juxtasomal labeling in S1 of a urethane-anesthetized rat, although we emphasize the general applicability of this technique to many preparations of choice.

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			<td>R. Bruno, Columbia Univ., NY, USA</td>
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			<td height="21" style="height:21px;width:224px;">Sugi absorbent swabs</td>
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			<td height="21" style="height:21px;width:224px;">Cytochrome C from equine heart</td>
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			<td height="21" style="height:21px;width:224px;">Vectastain standard ABC-kit</td>
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			<td style="width:275px;">Sigma</td>
			<td style="width:119px;">T9284</td>
			<td></td>
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			<td height="21" style="height:21px;width:224px;">Urethane</td>
			<td style="width:275px;">Sigma</td>
			<td style="width:119px;">U2500</td>
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			<td height="21" style="height:21px;width:224px;">Isoflurane</td>
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			<td style="width:119px;">45.112.110</td>
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			<td height="21" style="height:21px;width:224px;">Biocytin</td>
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			<td style="width:119px;">36219518</td>
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			<td height="21" style="height:21px;width:224px;">Analytical grade glycerol</td>
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			<td style="width:119px;">49767</td>
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			<td height="21" style="height:21px;width:224px;">HEPES</td>
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			<td>H3375</td>
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			<td height="21" style="height:21px;width:224px;">NaCl</td>
			<td style="width:275px;">Sigma Aldrich</td>
			<td style="width:119px;">31434</td>
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			<td height="21" style="height:21px;width:224px;">KCl</td>
			<td style="width:275px;">Sigma Aldrich</td>
			<td style="width:119px;">60130</td>
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			<td height="21" style="height:21px;width:224px;">CaCl</td>
			<td style="width:275px;">Sigma Aldrich</td>
			<td style="width:119px;">22,350-6</td>
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juxtasomal biocytin labeling to study the structure-function relationship of individual cortical neurons

The cerebral cortex is characterized by multiple layers and many distinct cell-types that together as a network are responsible for many higher cognitive functions including decision making, sensory-guided behavior or memory. To understand how such intricate neuronal networks perform such tasks, a crucial step is to determine the function (or electrical activity) of individual cell types within the network, preferentially when the animal is performing a relevant cognitive task. Additionally, it is equally important to determine the anatomical structure of the network and the morphological architecture of the individual neurons to allow reverse engineering the cortical network. Technical breakthroughs available today allow recording cellular activity in awake, behaving animals with the valuable option of post&#160;hoc identifying the recorded neurons. Here, we demonstrate the juxtasomal biocytin labeling technique, which involves recording action potential spiking in the extracellular (or loose-patch) configuration using conventional patch pipettes. The juxtasomal recording configuration is relatively stable and applicable across behavioral conditions, including anesthetized, sedated, awake head-fixed, and even in the freely moving animal. Thus, this method allows linking cell-type specific action potential spiking during animal behavior to reconstruction of the individual neurons and ultimately, the entire cortical microcircuit. In this video manuscript, we show how individual neurons in the juxtasomal configuration can be labeled with biocytin in the urethane-anaesthetized rat for post&#160;hoc identification and morphological reconstruction.

Detailed knowledge on 3D structure of individual neurons is crucial for elucidating organizational principles of neuronal networks. Our method involves a pipeline to achieve high quality biocytin labeling from an in vivo preparation, thereby allowing post&#160;hoc neuronal classification and detailed reconstruction of dendritic and axonal architecture of single neurons at high resolution. Depending on the quality of juxtasomal labeling, neurons are recovered with different DAB-intensities ranging from f...

Watch this Scientific Journal Video about Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons at JoVE.com

Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons

To understand the structure of neuronal networks, functional and morphological characterization of individual neurons is a necessity. Here, we demonstrate juxtasomal biocytin labeling, which allows electrophysiological recordings in the extracellular configuration, yet maintaining the ability to intracellularly label the neuron for post&#160;hoc reconstruction of dendritic and axonal architecture.

The cerebral cortex is characterized by multiple layers and many distinct cell-types that together as a network are responsible for many higher cognitive ...

juxtasomal-biocytin-labeling-to-study-structure-function-relationship

Research

JoVE Journal

Neuroscience

13.5K Views.  VU University Amsterdam. To understand the structure of neuronal networks, functional and morphological characterization of individual neurons is a necessity. Here, we demonstrate juxtasomal biocytin labeling, which allows electrophysiological recordings in the extracellular configuration, yet maintaining the ability to intracellularly label the neuron for post hoc reconstruction of dendritic and axonal architecture.

Video: Juxtasomal Biocytin Labeling to Study the Structure-function Relationship of Individual Cortical Neurons

Laser Capture Microdissection of Drosophila Peripheral Neurons

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the molecular mechanisms underlying class-specific dendrite morphogenesis1,2. To facilitate molecular analyses of class-specific da neuron development, it is vital to obtain these cells in a pure population. Although a range of different cell, and tissue-specific RNA isolation techniques exist for Drosophila cells, including magnetic bead based cell purification3,4, Fluorescent Activated Cell Sorting (FACS)5-8, and RNA binding protein based strategies9, none of these methods can be readily utilized for isolating single or multiple class-specific Drosophila da neurons with a high degree of spatial precision. Laser Capture Microdissection (LCM) has emerged as an extremely powerful tool that can be used to isolate specific cell types from tissue sections with a high degree of spatial resolution and accuracy. RNA obtained from isolated cells can then be used for analyses including qRT-PCR and microarray expression profiling within a given cell type10-16. To date, LCM has not been widely applied in the analysis of Drosophila tissues and cells17,18, including da neurons at the third instar larval stage of development. 
Here we present our optimized protocol for isolation of Drosophila da neurons using the infrared (IR) class of LCM. This method allows for the capture of single, class-specific or multiple da neurons with high specificity and spatial resolution. Age-matched third instar larvae expressing a UAS-mCD8::GFP19 transgene under the control of either the class IV da neuron specific ppk-GAL420 driver or the pan-da neuron specific 21-7-GAL421 driver were used for these experiments. RNA obtained from the isolated da neurons is of very high quality and can be directly used for downstream applications, including qRT-PCR or microarray analyses. Furthermore, this LCM protocol can be readily adapted to capture other Drosophila cell types a various stages of development dependent upon the cell type specific, GAL4-driven expression pattern of GFP.

Laser Capture Microdissection of Drosophila Peripheral Neurons

In this video-article we present a method for isolating single or multiple Drosophila da neurons from third instar larvae using the infrared capture (IR) class of Laser Capture Microdissection (LCM). RNA obtained from the isolated neurons can be readily used for downstream applications including qRT-PCR or microarray analyses.

The dendritic arborization (da) neurons of the Drosophila peripheral nervous system (PNS) provide an excellent model system in which to investigate the ...

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

DiI-Labeling of DRG Neurons to Study Axonal Branching in a Whole Mount Preparation of Mouse Embryonic Spinal Cord

Here we present a technique to label the trajectories of small groups of DRG neurons into the embryonic spinal cord by diffusive staining using the lipophilic tracer 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI)1. The comparison of axonal pathways of wild-type with those of mouse lines in which genes are mutated allows testing for a functional role of candidate proteins in the control of axonal branching which is an essential mechanism in the wiring of the nervous system. Axonal branching enables an individual neuron to connect with multiple targets, thereby providing the physical basis for the parallel processing of information. Ramifications at intermediate target regions of axonal growth may be distinguished from terminal arborization. Furthermore, different modes of axonal branch formation may be classified depending on whether branching results from the activities of the growth cone (splitting or delayed branching) or from the budding of collaterals from the axon shaft in a process called interstitial branching2 (Fig. 1).
The central projections of neurons from the DRG offer a useful experimental system to study both types of axonal branching: when their afferent axons reach the dorsal root entry zone (DREZ) of the spinal cord between embryonic days 10 to 13 (E10 - E13) they display a stereotyped pattern of T- or Y-shaped bifurcation. The two resulting daughter axons then proceed in rostral or caudal directions, respectively, at the dorsolateral margin of the cord and only after a waiting period collaterals sprout from these stem axons to penetrate the gray matter (interstitial branching) and project to relay neurons in specific laminae of the spinal cord where they further arborize (terminal branching)3. DiI tracings have revealed growth cones at the dorsal root entry zone of the spinal cord that appeared to be in the process of splitting suggesting that bifurcation is caused by splitting of the growth cone itself4 (Fig. 2), however, other options have been discussed as well5.
This video demonstrates first how to dissect the spinal cord of E12.5 mice leaving the DRG attached. Following fixation of the specimen tiny amounts of DiI are applied to DRG using glass needles pulled from capillary tubes. After an incubation step, the labeled spinal cord is mounted as an inverted open-book preparation to analyze individual axons using fluorescence microscopy.

The stereotyped projections of sensory afferents into the rodent spinal cord offer an easily accessible experimental system to study axonal branching through the tracing of single axons.

Here we  present a technique to label the trajectories of small groups of DRG neurons  into the embryonic spinal cord by diffusive staining using the ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

Differential Labeling of Cell-surface and Internalized Proteins after Antibody Feeding of Live Cultured Neurons

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of live neurons with a specific primary antibody raised against an extracellular portion of the protein. Given that receptors are trafficked to and from the surface, if cells are permeabilized after fixation then both cell-surface and internal protein will be detected by the same labeled secondary antibody. Here, we adapted a method used to study protein trafficking (&#8220;antibody feeding&#8221;) to differentially label protein that had been internalized by endocytosis during the antibody incubation step and protein that either remained on the cell surface or was trafficked to the surface during this period. The ability to distinguish these two pools of protein was made possible through the incorporation of an overnight blocking step with highly-concentrated unlabeled secondary antibody after an initial incubation of unpermeabilized neurons with a fluorescently-labeled secondary antibody. After the blocking step, permeabilization of the neurons allowed detection of the internalized pool with a fluorescent secondary antibody labeled with a different fluorophore. Using this technique we were able to obtain important information about the subcellular location of this putative receptor, revealing that it was, indeed, trafficked to the cell-surface in neurons. This technique is broadly applicable to a range of cell types and cell-surface proteins, providing a suitable antibody to an extracellular epitope is available.

We describe a method to label protein on the surface of living neurons using a specific polyclonal antibody to extracellular epitopes. Protein bound by the antibody on the cell surface and subsequently internalized via endocytosis can be distinguished from protein remaining on, or trafficked to, the surface during the incubation.

In order to demonstrate the cell-surface localization of a putative transmembrane receptor in cultured neurons, we labeled the protein on the surface of ...

Laser Capture Microdissection - A Demonstration of the Isolation of Individual Dopamine Neurons and the Entire Ventral Tegmental Area

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue sections on a microscope slide. When combined with immunohistochemistry, LCM can be used to isolate individual cells types based on a specific protein marker. Here, the LCM technique is described for collecting a specific population of dopamine neurons directly labeled with&#160;tyrosine hydroxylase immunohistochemistry and for isolation of the dopamine neuron containing region of the ventral tegmental area using indirect tyrosine hydroxylase immunohistochemistry on a section adjacent to those used for LCM. An infrared (IR) capture laser is used to both dissect individual neurons as well as the ventral tegmental area off glass slides and onto an LCM cap for analysis. Complete dehydration of the tissue with 100% ethanol and xylene is critical. The combination of the IR capture laser and the ultraviolet (UV) cutting laser is used to isolate individual dopamine neurons or the ventral tegmental area when using PEN membrane slides. A PEN membrane slide has significant advantages over a glass slide as it offers better consistency in capturing and collecting cells, is faster collecting large pieces of tissue, is less reliant on dehydration and results in complete removal of the tissue from the slide. Although removal of large areas of tissue from a glass slide is feasible, it is considerably more time consuming and frequently leaves some residual tissue behind. Data shown here demonstrate that RNA of sufficient quantity and quality can be obtained using these procedures for quantitative PCR measurements. Although RNA and DNA are the most commonly isolated molecules from tissue and cells collected with LCM, isolation and measurement of microRNA, protein and epigenetic changes in DNA can also benefit from the enhanced anatomical and cellular resolution obtained using LCM.

The isolation of individual dopamine neurons or the ventral tegmental area with direct or indirect immunohistochemistry is demonstrated using laser capture microdissection. Parameters for isolation of tissue from a glass slide using an infrared laser and from membrane slides using the combination of an infrared and ultraviolet laser are discussed.

Laser capture microdissection (LCM) is used to isolate a concentrated population of individual cells or precise anatomical regions of tissue from tissue ...

Immunohistochemistry and Multiple Labeling with Antibodies from the Same Host Species to Study Adult Hippocampal Neurogenesis

Adult neurogenesis is a highly regulated, multi-stage process in which new neurons are generated from an activated neural stem cell via increasingly committed intermediate progenitor subtypes. Each of these subtypes expresses a set of specific molecular markers that, together with specific morphological criteria, can be used for their identification. Typically, immunofluorescent techniques are applied involving subtype-specific antibodies in combination with exo- or endogenous proliferation markers. We herein describe immunolabeling methods for the detection and quantification of all stages of adult hippocampal neurogenesis. These comprise the application of thymidine analogs, transcardial perfusion, tissue processing, heat-induced epitope retrieval, ABC immunohistochemistry, multiple indirect immunofluorescence, confocal microscopy and cell quantification. Furthermore we present&#160;a sequential multiple immunofluorescence protocol which circumvents problems usually arising from the need of using primary antibodies raised in the same host species. It allows an accurate identification of all hippocampal progenitor subtypes together with a proliferation marker within a single section. These techniques are a powerful tool to study the regulation of different progenitor subtypes in parallel, their involvement in brain pathologies and their role in specific brain functions.

This video article illustrates a comprehensive protocol to detect and quantify all stages of adult hippocampal neurogenesis within the same tissue section. We elaborated a method to overcome the limitations of indirect multiple immunofluorescence that arise when suitable antibodies from different host species are unavailable.

Adult neurogenesis is a highly regulated, multi-stage process in which new neurons are generated from an activated neural stem cell via increasingly ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...