Supported by NIH R01 DC 17924, R01 DC 18401 (Klug), and NIH 1R15HD105231-01, T32DC012280, and FRAXA (McCullagh). The CARS imaging was performed in the Advanced Light Microscopy Core part of the NeuroTechnology Center at the University of Colorado Anschutz Medical Campus supported in part by NIH P30 NS048154 and NIH P30 DK116073.

All experiments complied with all applicable laws, National Institutes of Health guidelines, and were approved by the University of Colorado Anschutz Institutional Animal Care and Use Committee.
1. Animals
<ol>
	<li>Use C57BL/6J (stock #000664) mice (Mus musculus) obtained from The Jackson Laboratory or Mongolian gerbils (Meriones unguiculatus) originally obtained from Charles River.</li>
</ol>
2. Tissue preparation
<ol>
	<li>For transcardial perfusion, overdose rodent species of interest with pentobarbital (120 mg/kg body weight) and transcardially perfuse them with phosphate-buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 1.76 mM KH2PO4, 10 mM Na2HPO4) followed by 4% paraformaldehyde (PFA)24.
	<ol>
		<li>Specifically, open the abdomen and the rib cage using scissors and hold the rib cage in place with Kelly hemostatic forceps to expose the heart.</li>
		<li>Insert a 23 GA needle connected to a perfusion pump into the left ventricle and quickly cut the right atrium using fine scissors.</li>
		<li>Administer PBS through the perfusion pump and the needle in the heart for 10 min to clear the brain and body of blood.</li>
		<li>Switch the perfusion pump to 4% PFA for 10 min and check for rigidity of limbs and tail to confirm successful perfusion.</li>
	</ol>
	</li>
	<li>After perfusion, decapitate the animals and remove their brain from the skull. Keep the brains overnight in 4% PFA before transferring to PBS. Embed brainstems in 4% agarose (in PBS) and slice coronally using a vibratome at 200 &#181;m thickness.</li>
</ol>
3. Staining
<ol>
	<li>Stain free floating sections for Nissl, to visualize cell bodies (1:100), in antibody media (AB media: 0.1 M phosphate buffer (PB: 50 mM KH2PO4, 150 mM Na2HPO4), 150 mM NaCl, 3 mM Triton-X, 1% bovine serum albumin (BSA)) for 30 min at room temperature on a standard laboratory shaker25.
	<ol>
		<li>Protect sections from light using aluminum foil and/or a cover. 550 nm or below wavelengths are compatible with CARS imaging (Figure 1). 
		NOTE: While we do not expect that Triton-X or other reagents have an impact on CARS imaging of lipids, additional controls with specific antibody media may be warranted.</li>
	</ol>
	</li>
	<li>PAUSE POINT: Store free floating sections (while protected from light) in PBS until imaging. Once sectioned, image brain sections within 2 weeks.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64013/64013fig01.jpg" /> 
Figure 1: CARS imaging can be combined with immunofluorescent imaging. The graphs shows that CARS imaging occurs at 660/640 nm red signal spectrum26. This wavelength is sufficiently far removed from the green, blue, or UV range, allowing for combination of the CARS signal with immunofluorescence in these ranges. Specifically, the graph also indicates the excitation and emission for Nissl tagged with blue fluorophore, which was combined with CARS during the collection of representative results for this publication. <a href="https://www.jove.com/files/ftp_upload/64013/64013fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Imaging
NOTE: The CARS laser set up contains a fiber laser that provides the 80 MHz clock, and an OPO (Optical Parametric Oscillator) laser with a tunable range of 770-990 nm with the Stokes beam fixed at 1031 nm, which are needed for collecting the CARS signal. There is one aperture for both beams.
<ol>
	<li>Before bringing samples to the microscope, turn on and warm up CARS laser for at least 1 h, align the CARS laser, and Koehler the condenser optics and the diaphragm of the microscope for forward CARS imaging. 
	NOTE: This step is critical for proper function of CARS microscopy.
	<ol>
		<li>For spatial alignment of the two laser beams (pump and stokes), access the two internal PSDs (position sensitive detectors) via the CARS laser GUI.</li>
		<li>Achieve temporal alignment by using the delay function in the CARS laser GUI, which can help overlap the pulses of the two lasers (pump and stokes) that have different dispersions because of their different wavelengths. Therefore, both the temporal and spatial overlapping of the pump and stokes beams are done with the user input through the GUI.</li>
		<li>Adjust the external periscope (last two mirrors of the setup) to center the spatially overlapped two lasers onto the scanning head mirrors of the microscope.</li>
		<li>For best forward CARS non-descanned detection, make sure the condenser is Koehler-ed (meaning the condenser is centered and focused onto the diaphragm to achieve uniform illumination)</li>
	</ol>
	</li>
	<li>For immunofluorescence confocal imaging and CARS imaging, fit the CARS laser, with both forward and epi CARS non-descanned detectors (NDDs), by incorporating a confocal microscope equipped with visible lasers for fluorescence imaging (Figure 2).</li>
	<li>Place sections in a culture dish with coverslip (for inverted microscopy), PBS to avoid the tissue drying out, and a glass weight to keep tissue near coverslip.</li>
	<li>Take z-stacks or single images with a 60X, 1.2 NA infrared corrected water objective, which serves for collection of the CARS signal in the epi direction and through a 0.55 NA condenser in the forward direction for imaging brain areas such as the medial nucleus of the trapezoid body (MNTB).
	<ol>
		<li>Take the CARS images at approximately 600 mW pump/probe and 300 mW Stokes, by using the CARS laser GUI. These laser power values are measured internally by the system. Both lasers&#39; powers at the sample location are less than 25 mW and safe for the tissue sample.</li>
		<li>Overlap the pump and Stokes beams spatially and temporally. Tune the OPO to 797.2 nm. This yields a CARS wavelength of 650 nm. Because of the higher energy level, the resulting return to the ground state is anti-Stokes (blue shifted) to the excitation.</li>
		<li>Capture the CARS signal in epi or forward non-descanned detectors using bandpass filters (640-680 nm) followed by sequential detection of the immunofluorescence label (in this instance fluorescently labeled Nissl). 
		NOTE: The Nissl neuronal soma marker is not caught in the CARS 640-680 nm bandpass filter, allowing the combination of fluorescence and CARS imaging in the images presented below.</li>
		<li>The CARS and fluorescence do not share PMTs. Use these settings for optimal lipid signal to selectively image myelination in the brain area. 
		CAUTION: Shield the user from the laser beam</li>
	</ol>
	</li>
	<li>Save the images as .oib files, which can be imported into an image analysis program for further quantification (Figure 3).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64013/64013fig02.jpg" /> 
Figure 2: CARS instrument diagram showing CARS lasers (red arrows) and non-descanned (NDD) epi and forward detection incorporated onto a laser scanning confocal. In forward NDD we acquire CARS for C-H bonds (dark red arrows) and SHG (second harmonic generatioin) at 515 nm (orange arrow). In epi NDD, we acquire CARS for C-H bonds (dark red arrows) and 2PE (two-photon emission) autofluorescence (light blue arrow). Sequentially, fluorescence confocal images can be acquired (green arrows for visible laser, blue arrows for confocal detection). <a href="https://www.jove.com/files/ftp_upload/64013/64013fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64013/64013fig03v2.jpg" /> 
Figure 3: CARS can illuminate myelin (magenta) in brain tissue (brainstem) while also imaging Nissl (cyan) or fluorescent markers. The two panels show representative results from a Mongolian gerbil (single image M. unguiculatus, Figure 3A,C,E) and mouse (z-stack max projection M. musculus, Figure 2B,D,F) brain, indicating that this technique can be used across species. Figure 3A,B showing Nissl in cyan, C,D show the CARS signal in magenta, E,F combine the Nissl and CARS signals with each panel for gerbil or mouse, respectively. Both sets of images show a section of the medial nucleus of the trapezoid body (MNTB) in the brain stem. Neurons in the MNTB receive inputs from heavily myelinated axons, which terminate in the calyx of held, a type of giant synapse27. Scale bar is 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/64013/64013fig03largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors declare no conflicts of interest.

A growing body of literature emphasizes the role of myelin in brain function13,16,21,28. Moreover, we know that myelination thickness and myelination pattern can change in several neurological conditions such as multiple sclerosis (reviewed in29), aging (reviewed in30), autism20,31, and ma...

Action potentials are the basic unit of information in the brain, and action potential propagation through axons forms one pillar of information processing1,2,3. Neurons typically receive afferent inputs from multiple other neurons and integrate these inputs within a given narrow time window4,5. Therefore, the mechanisms of action potential propagation in axons have received a significant amount of attention from investigators.
When propagating through an axon, an action potential is regenerated repeatedly along the axon to ensure reliable propagation6. In most neurons of jawed vertebrates (gnathostomes) axons are surrounded by a sheath of myelin, which is a lipid-rich substance produced by nearby oligodendrocytes or Schwann cells, which are types of glial cells (reviewed in7,8). This myelin sheath electrically insulates the axon, reducing its capacitance and allowing action potential propagation efficiently, quickly, and with lower energy consumption. Myelin does not cover the axon uniformly, but it sheaths the axon in segments that have short gaps in between them, called the nodes of Ranvier (reviewed in9,10). Both myelination thickness, which controls the level of electrical insulation of an axon, and the spacing of the nodes of Ranvier, which control the frequency with which action potentials are regenerated along an axon, influence the speed of action potential propagation (reviewed in11).
There is a large body of literature suggesting that myelination thickness affects the speed of action potential propagation in axons12,13,14. Moreover, alterations in axon myelination can result in a number of CNS deficits15,16,17,18,19,20,21. It is therefore not surprising that the focus of many research efforts involves the measurement and characterization of axon myelination. Measurements of myelin thickness have most commonly been done with electron microscopy, a technique that requires a significant amount of tissue preparation and is challenging to use in combination with immunohistochemistry. However, there is also a faster and simpler technique to measure axon myelination that is based on Coherent Anti-Stokes Raman Spectroscopy (CARS). A CARS laser can be tuned to various frequencies and when tuned to frequencies that are suitable to excite lipids, myelin can be imaged without the need for any additional labels22. The lipid imaging can be combined with standard immunohistochemistry such that lipids can be imaged together with several fluorescent channels23. Imaging myelination with CARS is significantly faster than electron microscopy and has a resolution that is, albeit lower than EM, sufficient to detect even small differences in myelination in the same type of axons.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Anesthetic:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL disposable syringe with needle 27 GA x 0.5&#34;</td>
			<td>Exel int</td>
			<td>260040</td>
			<td></td>
		</tr>
		<tr>
			<td>Fatal +</td>
			<td>Vortech</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgery:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Spring Scissors - 8mm Cutting Edge</td>
			<td>Fine Science Tools</td>
			<td>15024-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Standard tweezers</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusion:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4% Paraformaldehyde</td>
			<td>Fisher Chemical</td>
			<td>SF994 (CS)</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors - Sharp</td>
			<td>Fine Science Tools</td>
			<td>14063-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Kelly hemostats</td>
			<td>Fine Science Tools</td>
			<td>13019-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Millipore H2O</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Needle tip, 23 GA x 1&#34;</td>
			<td>BD precision glide</td>
			<td>305193</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffered saline (PBS):</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Sigma</td>
			<td>P9333</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate monobase</td>
			<td>Sigma</td>
			<td>P5655</td>
			<td></td>
		</tr>
		<tr>
			<td>pump with variable flow or equivalent</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Fisher Chemical</td>
			<td>s271-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodiumphosphate dibasic</td>
			<td>Sigma</td>
			<td>S7907</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL vial with 4% PFA</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bochem Chemical Spoon 180mm</td>
			<td>Bochem</td>
			<td>230331000</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors - Sharp</td>
			<td>Fine Science Tools</td>
			<td>14063-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Noyes Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15011-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Pair of fine (Graefe) tweezers</td>
			<td>Fine Science Tools</td>
			<td>11050-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Shallow glass or plastic tray, approximately 10&#34; x 10&#34;</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Standard tweezers</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Scissors - Blunt</td>
			<td>Fine Science Tools</td>
			<td>14000-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Slicing:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Agar, plant</td>
			<td>RPI</td>
			<td>9002-18-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica</td>
			<td>VT1000s</td>
			<td></td>
		</tr>
		<tr>
			<td>well plate</td>
			<td>Alkali Sci.</td>
			<td>TPN1048-NT</td>
			<td></td>
		</tr>
		<tr>
			<td>Staining:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AB Media:</td>
			<td></td>
			<td></td>
			<td>1n 1,000 mL of Millipore H2O</td>
		</tr>
		<tr>
			<td>Phosphate buffered (PB):</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Phosphate Monobase</td>
			<td>Sigma</td>
			<td>P5655</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Phosohate Dibasic</td>
			<td>Sigma</td>
			<td>S7907</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA (Bovine serum albumin)</td>
			<td>Sigma life science</td>
			<td>A2153-100g</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>Fisher Chemical</td>
			<td>s271-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma - Aldrich</td>
			<td>x100-500ml</td>
			<td></td>
		</tr>
		<tr>
			<td>Nissl 435/455</td>
			<td>Invitrogen</td>
			<td>N21479</td>
			<td></td>
		</tr>
		<tr>
			<td>CARS:</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>APE&#160;picoemerald laser</td>
			<td>Angewandte Physik &#38; Elektronik&#160;GmbH</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>bandpass filter (420-520 nm)</td>
			<td>Chroma Technology</td>
			<td>HQ470/100m-2P</td>
			<td></td>
		</tr>
		<tr>
			<td>bandpass filter (500-530 nm)</td>
			<td>Chroma Technology</td>
			<td>HQ515/30m-2P</td>
			<td></td>
		</tr>
		<tr>
			<td>bandpass filters (640-680 nm)</td>
			<td>Chroma Technology</td>
			<td>HQ660/40m-2P</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope</td>
			<td>Olympus</td>
			<td>FV1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Cut Transfer pipet</td>
			<td>Fisher</td>
			<td>13-711-7M</td>
			<td></td>
		</tr>
		<tr>
			<td>dichroic longpass 565 nm</td>
			<td>Chroma Technology</td>
			<td>565dcxr</td>
			<td></td>
		</tr>
		<tr>
			<td>dichroic longpass 585 nm</td>
			<td>Chroma Technology</td>
			<td>585dcxr</td>
			<td></td>
		</tr>
		<tr>
			<td>dichroic shortpass 750 nm</td>
			<td>Chroma Technology</td>
			<td>T750spxrxt</td>
			<td></td>
		</tr>
		<tr>
			<td>glass bottom culture dish</td>
			<td>MatTek</td>
			<td>P35G-0-10-C</td>
			<td></td>
		</tr>
		<tr>
			<td>glass weight (10 mm x 10 mm boro rod)</td>
			<td>Allen Scientific Glass Inc</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>multiphoton shortpass emission filter 680 nm</td>
			<td>Chroma Technology</td>
			<td>ET680sp-2p8</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

coherent anti-stokes raman spectroscopy (cars) application for imaging myelination in brain slices

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature vibrations of molecules. However, these vibrational signatures are also characteristic of molecules within anatomical tissue such as the brain, making it increasingly useful and applicable for Neuroscience applications. For example, CARS can measure lipids by specifically exciting chemical bonds within these molecules, allowing for quantification of different aspects of tissue, such as myelin involved in neurotransmission. In addition, compared to other techniques typically used to quantify myelin, CARS can also be set up to be compatible with immunofluorescent techniques, allowing for co-labeling with other markers such as sodium channels or other components of synaptic transmission. Myelination changes are an inherently important mechanism in demyelinating diseases such as multiple sclerosis or other neurological conditions such as Fragile X Syndrome or autism spectrum disorders is an emerging area of research. In conclusion, CARS can be utilized in innovative ways to answer pressing questions in Neuroscience and provide evidence for underlying mechanisms related to many different neurological conditions.

One of the biggest advantages of CARS microscopy over other techniques is the compatibility with fluorescent imaging23. Figure 1 shows the CARS spectra compared to Nissl tagged with immunofluorescent marker showing little/no overlap in spectra. Figure 2 illustrates the laser set up for CARS in combination with confocal microscopy. Figure 3 demonstrates two representative images, one as a single stack and one ...

Watch this Scientific Journal Video about Coherent Anti-Stokes Raman Spectroscopy CARS Application for Imaging Myelination in Brain Slices at JoVE.com

Coherent Anti-Stokes Raman Spectroscopy CARS Application for Imaging Myelination in Brain Slices

Visualizing myelination is an important goal for many researchers studying the nervous system. CARS is a technique that is compatible with immunofluorescence that can natively image lipids within tissue such as the brain illuminating specialized structures such as myelin.

Coherent Anti-Stokes Raman Spectroscopy (CARS) Application for Imaging Myelination in Brain Slices

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature ...

coherent-anti-stokes-raman-spectroscopy-cars-application-for-imaging

Research

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Neuroscience

2.5K Views.  Oklahoma State University. Visualizing myelination is an important goal for many researchers studying the nervous system. CARS is a technique that is compatible with immunofluorescence that can natively image lipids within tissue such as the brain illuminating specialized structures such as myelin.

Video: Coherent Anti-Stokes Raman Spectroscopy CARS Application for Imaging Myelination in Brain Slices

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Imaging Calcium Responses in GFP-tagged Neurons of Hypothalamic Mouse Brain Slices

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the precise molecular steps as well as the activity of specific neurons in multi-functional nuclei of brain areas such as the hypothalamus remain. This problem includes identification of the molecular components involved in the regulation of various neurohormone signal transduction cascades. Elevations of intracellular Ca2+ play an important role in regulating the sensitivity of neurons, both at the level of signal transduction and at synaptic sites.
New tools have emerged to help identify neurons in the myriad of brain neurons by expressing green fluorescent protein (GFP) under the control of a particular promoter. To monitor both spatially and temporally stimulus-induced Ca2+ responses in GFP-tagged neurons, a non-green fluorescent Ca2+ indicator dye needs to be used. In addition, confocal microscopy is a favorite method of imaging individual neurons in tissue slices due to its ability to visualize neurons in distinct planes of depth within the tissue and to limit out-of-focus fluorescence. The ratiometric Ca2+ indicator fura-2 has been used in combination with GFP-tagged neurons1. However, the dye is excited by ultraviolet (UV) light. The cost of the laser and the limited optical penetration depth of UV light hindered its use in many laboratories. Moreover, GFP fluorescence may interfere with the fura-2 signals2. Therefore, we decided to use a red fluorescent Ca2+ indicator dye. The huge Stokes shift of fura-red permits multicolor analysis of the red fluorescence in combination with GFP using a single excitation wavelength. We had previously good results using fura-red in combination with GFP-tagged olfactory neurons3. The protocols for olfactory tissue slices seemed to work equally well in hypothalamic neurons4. Fura-red based Ca2+ imaging was also successfully combined with GFP-tagged pancreatic &beta;-cells and GFP-tagged receptors expressed in HEK cells5,6. A little quirk of fura-red is that its fluorescence intensity at 650 nm decreases once the indicator binds calcium7. Therefore, the fluorescence of resting neurons with low Ca2+ concentration has relatively high intensity. It should be noted, that other red Ca2+-indicator dyes exist or are currently being developed, that might give better or improved results in different neurons and brain areas.

In this protocol, we update recent progress in imaging Ca2+ signals of GFP-tagged neurons in brain tissue slices using a red fluorescent Ca2+ indicator dye.

Despite an enormous increase in our knowledge about the mechanisms underlying the encoding of information in the brain, a central question concerning the ...

Local Application of Drugs to Study Nicotinic Acetylcholine Receptor Function in Mouse Brain Slices

Tobacco use leads to numerous health problems, including cancer, heart disease, emphysema, and stroke. Addiction to cigarette smoking is a prevalent neuropsychiatric disorder that stems from the biophysical and cellular actions of nicotine on nicotinic acetylcholine receptors (nAChRs) throughout the central nervous system. Understanding the various nAChR subtypes that exist in brain areas relevant to nicotine addiction is a major priority.
Experiments that employ electrophysiology techniques such as whole-cell patch clamp or two-electrode voltage clamp recordings are useful for pharmacological characterization of nAChRs of interest. Cells expressing nAChRs, such as mammalian tissue culture cells or Xenopus laevis oocytes, are physically isolated and are therefore easily studied using the tools of modern pharmacology. Much progress has been made using these techniques, particularly when the target receptor was already known and ectopic expression was easily achieved. Often, however, it is necessary to study nAChRs in their native environment: in neurons within brain slices acutely harvested from laboratory mice or rats. For example, mice expressing "hypersensitive" nAChR subunits such as &alpha;4 L9&prime;A mice 1 and &alpha;6 L9&prime;S mice 2, allow for unambiguous identification of neurons based on their functional expression of a specific nAChR subunit. Although whole-cell patch clamp recordings from neurons in brain slices is routinely done by the skilled electrophysiologist, it is challenging to locally apply drugs such as acetylcholine or nicotine to the recorded cell within a brain slice. Dilution of drugs into the superfusate (bath application) is not rapidly reversible, and U-tube systems are not easily adapted to work with brain slices.
In this paper, we describe a method for rapidly applying nAChR-activating drugs to neurons recorded in adult mouse brain slices. Standard whole-cell recordings are made from neurons in slices, and a second micropipette filled with a drug of interest is maneuvered into position near the recorded cell. An injection of pressurized air or inert nitrogen into the drug-filled pipette causes a small amount of drug solution to be ejected from the pipette onto the recorded cell. Using this method, nAChR-mediated currents are able to be resolved with millisecond accuracy. Drug application times can easily be varied, and the drug-filled pipette can be retracted and replaced with a new pipette, allowing for concentration-response curves to be created for a single neuron. Although described in the context of nAChR neurobiology, this technique should be useful for studying many types of ligand-gated ion channels or receptors in neurons from brain slices.

In this paper, we describe a useful method to study ligand-gated ion channel function in neurons of acutely isolated brain slices. This method involves the use of a drug-filled micropipette for local application of drugs to neurons recorded using standard patch clamp techniques.

Tobacco use leads to numerous health problems, including cancer, heart disease, emphysema, and stroke. Addiction to cigarette smoking is a prevalent ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Paired Whole Cell Recordings in Organotypic Hippocampal Slices

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.

Pair recordings are simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling precise electrophysiological and pharmacological characterization of the synapses between individual neurons. Here we describe the detailed methodology and requirements for establishing this technique in organotypic hippocampal slice cultures in any laboratory equipped for electrophysiology.

Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a challenging and low-throughput technique due to its complexity and high reliance on user operation and control. This manuscript demonstrates an image-guided automatic patch clamp system for in vitro whole-cell patch clamp experiments in acute brain slices. Our system implements a computer vision-based algorithm to detect fluorescently labeled cells and to target them for fully automatic patching using a micromanipulator and internal pipette pressure control. The entire process is highly automated, with minimal requirements for human intervention. Real-time experimental information, including electrical resistance and internal pipette pressure, are documented electronically for future analysis and for optimization to different cell types. Although our system is described in the context of acute brain slice recordings, it can also be applied to the automated image-guided patch clamp of dissociated neurons, organotypic slice cultures, and other non-neuronal cell types.

This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a ...

Whole-cell Currents Induced by Puff Application of GABA in Brain Slices

Pharmacological administration is commonly used when conducting whole-cell patch-clamp recording in brain slices. One of the best methods of drug application during electrophysiological recording is the puff technique, which can be used to study the effect of pharmacological reagents on neuronal activities in brain slices. The greatest advantage of puff application is that the drug concentration around the recording site increases rapidly, thus preventing desensitization of membrane receptors. Successful use of puff application involves careful attention to the following elements: the concentration of the drug, the parameters of the puff micropipette, the distance between the tip of a puff micropipette and the neuron recorded, and the duration and pressure driving the puff (pounds per square inch, psi). This article describes a step-by-step procedure for recording whole-cell currents induced by puffing gamma-aminobutyric acid (GABA) onto a neuron of a prefrontal cortical slice. Notably, the same procedure can be applied with minor modifications to other brain areas such as the hippocampus and the striatum, and to different preparations, such as cell cultures.

We describe the puff technique, by which pharmacological reagents can be administered during whole-cell patch-clamp recording, and highlight various aspects of the features that are crucial for its success.