We thank Yehui Qin and Mallory Myers for technical support. We also thank Dr. Sigismund Huck for providing us the anti-&#945;4-ECD antibody. This work is supported by National Institutes of Health grant NS22061 to L. W. R.

All animal experiments were carried out in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publications No. 80-23, revised 2012) and studies were approved by Institutional Animal Care and Use for Research Committees at Stony Brook University (#1618 and #1792).
1. vHipp-nAcc Synaptic Co-cultures
<ol>
	<li>Sacrifice mice (postnatal day 0 - 3, from wild-type (WT) or &#945;7 nAChRs transgenic mouse line) with CO2. Decapitate each pup and save the tail in a 1.5 ml centrifuge tube for post facto genotyping. Perform the following steps in a sterilized hood.</li>
	<li>Remove the top of the skull with scissors and forceps to expose the whole brain. Starting at the juncture where the cerebral cortices part from each other caudally, obtain a coronal section that contains the posterior cortices including the vHipp5,21 under a dissecting microscope.</li>
	<li>Dissect vHipp tissue strips containing the CA1-subiculum region out according to a developing mouse brain atlas22 (Figure 1A). Transfer to a 35 mm culture dish containing cold culture media (4 &#176;C), cut into small thin slices (around 100 &#181;m &#215; 100 &#181;m microslices).</li>
	<li>Pre-incubate 12 mm poly-D-lysine/laminin-coated glass coverslips inside 24 well tissue culture plates with culture media (~ 500 &#956;l) for at least 30 min in the incubator at 37 &#176;C. Remove media before plating microslices.</li>
	<li>Plate with a fire-polished Pasteur pipette at the center of the coverslip with minimal amount of media (~ 50 &#956;l, containing ~ 20 pieces of microslices) to facilitate attachment of the explants. Plate vHipp microslices originating from a single animal (either the +/+ or the -/- genotype of the transgenic &#945;7 line) on each coverslip.</li>
	<li>After the explants settle onto the coverslip, gently add additional media (~ 100 &#956;l) by the wall so that the level of the media is high enough to completely cover the explants on the periphery, but not those at the center of the coverslip.</li>
	<li>After an O/N incubation in 37 &#176;C, 5% CO2 incubator, disperse nAcc neurons from embryonic days 18 to postnatal day 1 WT mice and add to the explants.
	<ol>
		<li>Dissect out nAcc tissues according to a developing mouse brain atlas20 (Figure 1B).</li>
		<li>Cut into small pieces (around 500 &#181;m &#215; 500 &#181;m microslices) and transfer to a 15 ml tube.</li>
		<li>Treat with 0.25% trypsin (2 ml) for 15 min at 37 &#176;C. Wash tissue chunks three times with cold washing media (5 ml, 4 &#176;C) followed by one wash in cold culture media (5 ml, 4 &#176;C). Allow suspension to rest for 5 min in each wash step, and then pour off supernatant carefully.</li>
		<li>Dissociate cells by gentle trituration in 2 ml of culture media with a lightly fire-polished Pasteur pipette.</li>
		<li>Transfer cell suspension to another 15 ml tube and centrifuge at 2,000 x rpm for 5 min. Remove the supernatant and resuspend the cells in culture media at 1 ml per 6 pups. Disperse cells by gently pipetting in media several times with a fire-polished Pasteur pipette.</li>
		<li>Add 0.25 ml of dispersed nAcc cells to each coverslip with vHipp explants.</li>
		<li>Maintain the cultures in a humidified 37 &#176;C, 5% CO2 incubator. Place culture plates on dampened sterile gauze pads to mechanically stabilize and maintain humidity, facilitating synapse formation.</li>
	</ol>
	</li>
</ol>
2. Immunocytochemistry
<ol>
	<li>Fix cultures (5 - 7 days in vitro) in 4% paraformaldehyde/4% sucrose/PBS (~500 &#956;l per coverslip, 20 min, RT).</li>
	<li>Permeabilize cultures with 0.25% Triton X-100/ PBS (~500 &#956;l per coverslip, 5 min, RT).</li>
	<li>Block cultures with 10% normal donkey serum in PBS (~500 &#956;l per coverslip, 30 min, RT) prior to antibody staining. Incubate with primary antibodies O/N at 4 &#176;C. Use the following primary antibodies: anti- nAChRs (&#945;4-ECD 1:500; anti-&#945;5 subunits, 1:500), anti-vesicular glutamate transporter 1 (1:250), anti-GAD65 (1:100).</li>
	<li>Wash primary antibodies out with PBS (3 times/~500 &#956;l per coverslip, 5 min) and then incubate cultures in secondary antibodies conjugated to Alexa 488 (~500 &#956;l per coverslip, 1:500) or Alexa 594 (~500 &#956;l per coverslip, 1:500) for 1 hr at 37 &#176;C.</li>
	<li>Incubate cultures in &#945;BgTx conjugated to Alexa 594 (~500 &#956;l per coverslip, 1:1,000 in culture media) for 45 min at 37 &#176;C prior to fixation for labeling surface &#945;7*nAChRs.</li>
	<li>Mount and seal coverslips.</li>
	<li>Capture images with a fluorescence microscope equipped with Plan-Apochromat objectives (20X with 0.8 NA or 63X oil with 1.4 NA) and a CCD camera.</li>
</ol>
3. FM1-43 Based Vesicular Fusion
<ol>
	<li>Maintain cultures (5 - 7 days in vitro) in an imaging chamber, mount the chamber on spinning disk confocal microscope. Continuously perfuse (1 ml/min) with HBS cocktail at RT.</li>
	<li>Stop perfusion and load cultures with 10 &#181;M FM1-43 in 56 mM K+ ACSF (~500 &#956;l) for 90 sec (staining), wash external dye away for 15 min with Ca2+ free HBS containing ADVASEP-7 (0.1 mM) to scavenge membrane-bound FM1-43.</li>
	<li>Stimulate the cultures with 56 mM K+ ACSF (~500 &#956;l) without FM1-43 for 120 sec (destaining), reload cultures with FM1-43 using the same conditions, and wash again with Ca2+ free HBS containing ADVASEP-7 for 15 min.</li>
	<li>Acquire FM1-43 fluorescence images with spinning disk confocal microscope equipped with a Plan-Apochromat objective (60X water with 1.4 NA, excitation 488 nm, emission 530 nm) and capture images with a CCD camera.</li>
	<li>Collect baseline FM1-43 fluorescence images (every 2 s for 1 min) as pre-nicotine control; apply nicotine (1 &#181;M) by rapid perfusion (2 ml/min) for 1 min, and then wash nicotine out with HBS cocktail. Keep capture time-lapse images before, during, and after nicotine application every 2 sec for 5 min.</li>
	<li>Quantify FM1-43 fluorescence intensity before (Fstaining) and after (Fdestaining) nicotine application at each synaptic bouton.</li>
	<li>Calculate total amount of releasable FM1-43 fluorescence along vHipp axons by quantifying the difference of FM1-43 fluorescence intensity before and after nicotine application (&#916;F = Fstaining-Fdestaining). Analyze fraction of fluorescence intensity decrease after nicotine with the following equation: Fdecrease% = &#916;F/Fstaining.</li>
</ol>
4. Calcium Imaging
<ol>
	<li>Rinse cultures (5 - 7 days in vitro) quickly with normal HBS, then load cultures with 5 &#181;M Fluo-4 Ca2+ indicator (AM ester) and 0.02% Pluronic F-127 in HBS (2 ml) for 30 min at 37 &#176;C and 5% CO2.</li>
	<li>Wash out Fluo-4 solution with HBS (3 times/5 min). Return cultures to incubator (37 &#176;C / 5% CO2) for at least 30 min.</li>
	<li>Maintain cultures in the same imaging chamber and same conditions as described in FM1-43 based vesicular fusion section (see step 3.1).</li>
	<li>Collect Fluo-4 fluorescence images of axonal projections from the vHipp micro-slices with a Plan-Apochromat objective (60X water with 1.4 NA, excitation 488 nm, emission 530 nm) and a CCD camera.</li>
	<li>Collect baseline Fluo-4 fluorescence images (every 10 sec for 2 min) as pre-nicotine control, apply nicotine (1 &#181;M) by rapid perfusion (2 ml/min) for 1 min, and then wash nicotine out with HBS cocktail. Keep capture time-lapse images before, during, and after nicotine application every 10 sec for 30 min.</li>
	<li>Save all frames of the raw fluo-4 fluorescence images, export as a series of TIFF format images for further analyzing.</li>
	<li>Collect the integrated intensity of fluo-4 fluorescence along vHipp axons before and after nicotine application.</li>
	<li>Normalize integrated fluorescence intensity with the following equation: &#916;F/F0 =&#160; (F-F0)/F0, where F0 is the background-corrected pre-nicotine integrated fluorescence intensity and F is the integrated fluorescence intensity along vHipp axons at each time point after nicotine application. Analyze and plot normalized integrated fluorescence intensity.</li>
</ol>

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The authors have nothing to disclose.&#160;

The co-culture preparation described re-capitulates ventral hippocampal&#8211;accumbens circuits in vitro. This preparation permits relatively straightforward and reliable examination of the spatial and temporal profiles by which activation of pre-synaptic nAChRs elicit enhanced glutamatergic transmission5, 21.
Co-cultures are defined as the growth of different specific cell types in one dish which can provide physiological conditions in vitro to demonstrate in...

Cholinergic modulation of circuit excitability contributes to fundamental aspects of cognition, and altered cholinergic modulation is a feature of neurodegenerative and neuropsychiatric disorders including Alzheimer&#8217;s disease, Parkinson&#8217;s disease, schizophrenia and addiction1-4. An established mechanism of cholinergic facilitation of synaptic transmission in the CNS is via direct activation of nAChRs localized at pre-synaptic sites. Activation of these pre-synaptic receptors leads to increased intracellular Ca2+ ([Ca2+]i) in pre-synaptic terminals &#8211; both directly, due to the relatively high calcium conductance of certain nAChR subtypes, and indirectly, via intracellular signaling cascades5, thereby enhancing neurotransmitter release. In fact, the activation of pre-synaptic nAChRs has been linked with changes in release of a wide variety of neurotransmitters including glutamate, GABA, ACh, and dopamine6-10. Although this process has been studied indirectly using electrophysiological methods at various synapses, optical reporters of [Ca2+]i and synaptic vesicle recycling allow more direct and temporally precise measurement of pre-synaptic phenomena.
Pre-synaptic localization of nAChRs has been demonstrated convincingly with direct immuno-gold labeling of nAChRs at the electron microscopic (EM) level11,12. Several other techniques have also been used to address the nAChR localization indirectly, including detecting locations of nAChRs subunit- fluorescent protein chimeras in cultured neurons13,14, electrophysiological recording of nAChR currents in synaptic terminals15,16, monitoring nicotine induced changes in [Ca2+]i in synaptic nerve terminals by live cell imaging17, and indirect monitoring of neurotransmitter release at the synaptic terminal by live cell imaging techniques with fluorescent indicators, including exocytosis of synaptic vesicles viewed by styryl amphipathic FM dyes (FM1-43 and FM4-64) and/or synapto-pHluorin and by specific fluorescent neurotransmitter reporters, such as CNiFERs for ACh and iGluSnFr for glutamate18-20. Overall, these current approaches for identifying pre-synaptic localization of nAChRs are complicated, and require special systems and techniques to allow reliable identification and physiological monitoring of pre-synaptic activity.
Here we describe protocols and equipment for an in vitro co-culture system of a ventral hippocampal (vHipp) &#8211; nucleus accumbens (nAcc) circuit that provides direct access to identify and analyze both pre- and post-synaptic components of synaptic transmission. We show examples of pre-synaptic localization of nAChRs and the live cell imaging of nAChR mediated Ca2+ signaling and neurotransmitter release along vHipp axons. A natural (and straightforward) extension of the protocol presented here is the preparation of pre- and post-synaptic contacts comprised of neurons from different genotypes. In this manner the contribution of a particular gene product to the pre- and/or post-synaptic mechanisms of modulation can be assessed directly.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>1, Culture Media (50 ml)</td>
		</tr>
		<tr>
			<td>Neurobasal&#160;</td>
			<td>GIBCO</td>
			<td>10888022</td>
			<td>48 ml</td>
		</tr>
		<tr>
			<td>B-27 Supplements</td>
			<td>GIBCO</td>
			<td>0080085-SA</td>
			<td>1 ml</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>GIBCO</td>
			<td>10908-010</td>
			<td>0.5 ml</td>
		</tr>
		<tr>
			<td>GlutaMAX Supplement</td>
			<td>GIBCO</td>
			<td>35050-061</td>
			<td>0.5 ml</td>
		</tr>
		<tr>
			<td>Brain-derived neurotrophic factor (BDNF)</td>
			<td>GIBCO</td>
			<td>15140-122</td>
			<td>20 ng/ml</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>2, washing media (HBSS, 100 ml)</td>
		</tr>
		<tr>
			<td>HBSS, no calcium, no magnesium, no phenol red&#160;</td>
			<td>GIBCO</td>
			<td>14175-095</td>
			<td>99 ml</td>
		</tr>
		<tr>
			<td>HEPES ( 1M)</td>
			<td>GIBCO</td>
			<td>15630-130</td>
			<td>1 ml</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>3, HEPES buffered saline&#160; (HBS)&#160;&#160; pH=7.3</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888&#160;</td>
			<td>135 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333&#160;</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266&#160;</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>CaCl2,</td>
			<td>Sigma</td>
			<td>C1016&#160;</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>4, HBS Cocktail for live imaging pH=7.3</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888&#160;</td>
			<td>135 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333&#160;</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266&#160;</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>CaCl2,</td>
			<td>Sigma</td>
			<td>C1016&#160;</td>
			<td>2 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>tetrodotoxin&#160;</td>
			<td>Tocris</td>
			<td>1078</td>
			<td>2 &#181;M</td>
		</tr>
		<tr>
			<td>bicuculline</td>
			<td>Tocris</td>
			<td>131</td>
			<td>10 &#181;M</td>
		</tr>
		<tr>
			<td>D-AP-5</td>
			<td>Tocris</td>
			<td>105</td>
			<td>50 &#181;M</td>
		</tr>
		<tr>
			<td>CNQX</td>
			<td>Tocris</td>
			<td>1045</td>
			<td>20 &#181;M</td>
		</tr>
		<tr>
			<td>LY341495</td>
			<td>Tocris</td>
			<td>1209</td>
			<td>10 &#181;M</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>5, Calcium-free&#160; HBS&#160;&#160; pH=7.3</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888&#160;</td>
			<td>135 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333&#160;</td>
			<td>5 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M8266&#160;</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>6, 56 mM Potassium ACSF pH=7.4</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S9888</td>
			<td>119 mM</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9333</td>
			<td>56 mM</td>
		</tr>
		<tr>
			<td>MgSO4.7H</td>
			<td>Sigma</td>
			<td>M1880</td>
			<td>1.3 mM</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma</td>
			<td>C1016</td>
			<td>2.5 mM</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Sigma</td>
			<td>S8282</td>
			<td>1 mM</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma</td>
			<td>S5761</td>
			<td>26.2 mM</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Sigma</td>
			<td>G0350500&#160;</td>
			<td>10 mM</td>
		</tr>
	</tbody>
</table>

live imaging of nicotine induced calcium signaling and neurotransmitter release along ventral hippocampal axons

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors (nAChRs) is an important mechanism for neuromodulation by acetylcholine (ACh). The difficulty with access to probing the signaling mechanisms within intact axons and at nerve terminals both in vitro and in vivo has limited progress in the study of the pre-synaptic components of synaptic plasticity. Here we introduce a gene-chimeric preparation of ventral hippocampal (vHipp)&#8211;accumbens (nAcc) circuit in vitro that allows direct live imaging to analyze both the pre- and post-synaptic components of transmission while selectively varying the genetic profile of the pre- vs post-synaptic neurons. We demonstrate that projections from vHipp microslices, as pre-synaptic axonal input, form multiple, reliable glutamatergic synapses with post-synaptic targets, the dispersed neurons from nAcc. The pre-synaptic localization of various subtypes of nAChRs are detected and the pre-synaptic nicotinic signaling mediated synaptic transmission are monitored by concurrent electrophysiological recording and live cell imaging. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of glutamatergic synaptic plasticity in vitro.

The preparation employed consists of gene chimeric co-cultures of vHipp&#8211;nAcc circuits in vitro. Projections emanating from vHipp microslices, as pre-synaptic axonal input, can make synaptic contacts with post-synaptic targets, the dispersed neurons from nAcc. Nicotine induced a sustained (&#8805; 30 min) facilitation of glutamatergic transmission from nAcc neurons innervated by vHipp axons21 and prolonged calcium signaling along vHipp axons5 via pre-synaptic &#945;7*nAChRs.
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Watch this Scientific Journal Video about Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons at JoVE.com

Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons

We developed a gene-chimeric preparation of ventral hippocampal &#8211; accumbens circuit in vitro that allows direct live imaging to analyze presynaptic mechanisms of nicotinic acetylcholine receptors (nAChRs) mediated synaptic transmission. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of synaptic plasticity.

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors ...

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8.9K visualizações.  Stony Brook University. We developed a gene-chimeric preparation of ventral hippocampal – accumbens circuit in vitro that allows direct live imaging to analyze presynaptic mechanisms of nicotinic acetylcholine receptors (nAChRs) mediated synaptic transmission. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of synaptic plasticity.

Vídeo: Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Analyzing Murine Schwann Cell Development Along Growing Axons

The development of peripheral nerves is an intriguing process. Neurons send out axons to innervate specific targets, which in humans are often more than 100 cm away from the soma of the neuron. Neuronal survival during development depends on target-derived growth factors but also on the support of Schwann cells (SCs). To this end SC ensheath axons from the region of the neuronal soma (or the transition from central to peripheral nervous system) to the synapse or neuromuscular junction. Schwann cells are derivatives of the neural crest and migrate as precursors along emerging axons until the entire axon is covered with SCs. This shows the importance of SC migration for the development of the peripheral nervous system and underlines the necessity to investigate this process. In order to analyze SC development, a setup is needed which next to the SCs also includes their physiological substrate for migration, the axon. Due to intrauterine development in vivo time-lapse imaging, however, is not feasible in placental vertebrates like mouse (mus musculus). To circumvent this, we adapted the superior cervical ganglion (SCG) explant technique. Upon treatment with nerve growth factor (NGF) SCG explants extend axons, followed by SC precursors migrating along the axons from the ganglion to the periphery. The beauty of this system is that the SC are derived from a pool of endogenous SC and that they migrate along their own physiological axons which are growing at the same time. This system is especially intriguing, because the SC development along axons can be analyzed by time-lapse imaging, opening further possibilities to gain insights into SC migration.

Here we describe a Schwann cell (SC) migration assay in which SCs are able to develop along extending axons.

The  development of peripheral nerves is an intriguing process. Neurons send out  axons to innervate specific targets, which in humans are often more than ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Dorsal Root Ganglia Isolation and Primary Culture to Study Neurotransmitter Release

Dorsal root ganglia (DRG) contain cell bodies of sensory neurons. This type of neuron is pseudo-unipolar, with two axons that innervate peripheral tissues, such as skin, muscle and visceral organs, as well as the spinal dorsal horn of the central nervous system. Sensory neurons transmit somatic sensation, including touch, pain, thermal, and proprioceptive sensations. Therefore, DRG primary cultures are widely used to study the cellular mechanisms of nociception, physiological functions of sensory neurons, and neural development. The cultured neurons can be applied in studies involving electrophysiology, signal transduction, neurotransmitter release, or calcium imaging. With DRG primary cultures, scientists may culture dissociated DRG neurons to monitor biochemical changes in single or multiple cells, overcoming many of the limitations associated with in vivo experiments. Compared to commercially available DRG-hybridoma cell lines or immortalized DRG neuronal cell lines, the composition and properties of the primary cells are much more similar to sensory neurons in tissue. However, due to the limited number of cultured DRG primary cells that can be isolated from a single animal, it is difficult to perform high-throughput screens for drug targeting studies. In the current article, procedures for DRG collection and culture are described. In addition, we demonstrate the treatment of cultured DRG cells with an agonist of neuropeptide FF receptor type 2 (NPFFR2) to induce the release of peptide neurotransmitters (calcitonin gene-related peptide (CRGP) and substance P (SP)).

Dorsal root ganglia (DRG) primary cultures are frequently used to study physiological functions or pathology-related events in sensory neurons. Here, we demonstrate the use of lumbar DRG cultures to detect the release of neurotransmitters after neuropeptide FF receptor type 2 stimulation with a selective agonist.

Dorsal root ganglia (DRG) contain cell bodies of sensory neurons. This type of neuron is pseudo-unipolar, with two axons that innervate peripheral ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its highly-organized subcellular architecture, we recently developed a new imaging modality, named spectral reflectometry (SpeRe), which enables unprecedented label-free nanoscale imaging of the live myelinated axons in situ. The underlying principle is to obtain nanostructural information by analyzing the reflectance spectrum of the multilayered subcellular structure. In this article, we describe a detailed step-by-step protocol for performing a basic SpeRe imaging of the nervous tissues using a commercial confocal microscopic system, equipped with a white-light laser and a tunable filter. We cover the procedures of sample preparation, acquisition of spectral data, and image processing for obtaining nanostructural information.

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its ...

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of individual cells. Since an increase of intracellular calcium (Ca2+) in the cytosol often occurs because of the electrical activity, or in response to a myriad of other stimuli, this process can be monitored by the imaging of cells loaded with fluorescent calcium-sensitive dyes.&#160; However, it is difficult to image this response in an individual cell type within whole tissue because these dyes are taken up by all cell types within the tissue. In contrast, genetically encoded calcium indicators (GECIs) can be expressed by an individual cell type and fluoresce in response to an increase of intracellular Ca2+, thus permitting the imaging of Ca2+ signaling in entire populations of individual cell types. Here, we apply the use of the GECIs GCaMP3/6 to the mouse neuromuscular junction, a tripartite synapse between motor neurons, skeletal muscle, and terminal/perisynaptic Schwann cells. We demonstrate the utility of this technique in classic ex vivo tissue preparations. Using an optical splitter, we perform dual-wavelength imaging of dynamic Ca2+ signals and a static label of the neuromuscular junction (NMJ) in an approach that could be easily adapted to monitor two cell-specific GECI or genetically encoded voltage indicators (GEVI) simultaneously. Finally, we discuss the routines used to capture spatial maps of fluorescence intensity. Together, these optical, transgenic, and analytic techniques can be employed to study the biological activity of distinct cell subpopulations at the NMJ in a wide variety of contexts.

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

Here we present a protocol to image calcium signaling in populations of individual cell types at the murine neuromuscular junction.

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of ...

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.