We especially thank Timo Fritsch for excellent animal care and Hermann D&#246;ring, Mohamed Elsaey, Sol Pose-M&#233;ndez, Jakob von Trotha, Komali Valishetti and Barbara Winter for their helpful support. We are also grateful to all the other members of the K&#246;ster lab for their feedback. The project was funded in part by the German Research Foundation (DFG, KO1949/7-2) project 241961032 (to RWK) and the Bundesministerium f&#252;r Bildung und Forschung (BMBF; Era-Net NEURON II CIPRESS project 01EW1520 to JCM) is acknowledged.

All animal work described here is in accordance with legal regulations (EU-Directive 2010/63). Maintenance and handling of fish have been approved by local authorities and by the animal welfare representative of the Technische Universit&#228;t Braunschweig.
1. Preparation of artificial cerebro spinal fluid (ACSF), low melting agarose and sharp glass needles
<ol>
	<li>Prepare the ACSF by dissolving the listed chemicals at following concentrations in distilled water. 134 mM NaCl (58.44 g/mol), 2.9 mM KCl (74.55 g/mol), 2.1 mM CaCl2 (110.99 g/mol), 1.2 mM MgCl2 6x H2O (203.3 g/mol), 10 mM HEPES (238.31 g/mol), and 10 mM d-Glucose (180.16 g/mol). 
	NOTE: For MgCl2, CaCl2, and KCl, 1 M stock solutions are prepared in desalted sterile water and stored at 4&#160;&#176;C for subsequently preparing fresh ACSF. Glucose, HEPES, and NaCl are dissolved as solid compounds in the fresh ACSF solution. For dissolving chemicals, follow the manufacturer&#39;s instructions.</li>
	<li>Adjust the pH of the ACSF to 7.8 with 10 M NaOH. Preparation of ACSF requires precise measurement of chemicals and fine adjustment of pH as it replaces the cerebro spinal fluid and maintains the physiological conditions required for neurons to be fully functional else it might cause brain misfunction and neuronal death.</li>
	<li>Store the freshly prepared ACSF at 4 &#176;C for a maximum of 4 weeks. For working conditions, aliquot the required volume of ACSF for the day/experiment and prewarm at 25-28 &#176;C (and optionally oxygenate it, step 2.5) 
	NOTE: Freshly prepared ASCF is fine for 1 day. If planning to use it over several days, ACSF needs to be sterile filtered.</li>
	<li>For later anesthesia of the larvae, prepare a 50 mM stock solution of d-Tubocurarine in distilled water and store the solution at -20 &#176;C as 100 &#181;L aliquots in the freezer until needed.</li>
	<li>To embed the fish, prepare 2.5% low melting (LM) agarose by dissolving 1.25 g LM-agarose (Table of Materials) in 50 mL ACSF and boil until the agarose is completely dissolved. 
	NOTE: Alternatively, higher or lower concentrations of LM-agarose can be used depending on the experimental set-up. However, if the agarose is too soft, it will not be able to hold the fish in position when opening the skull.</li>
	<li>Store the agarose at 37 &#176;C waterbath, to avoid solidification and because this temperature will also not harm the larvae when embedding. After the boiled agarose is cooled down to 37 &#176;C in the waterbath, add the necessary amount of d-Tubocurarine to the aliquoted agarose needed for the day to reach a working concentration of 10 &#181;M. For future use, store the left-over agarose at 4 &#176;C to avoid contamination.</li>
	<li>Prepare sharp and thin glass needles from glass capillaries (Supplementary Figure 1) using a micropipette puller with the following settings.
	<ol>
		<li>Puller I, Capillary type 1: Heat 1: 65.8; Heat 2: 55.1; 2-step pulling 
		Puller II, Capillary type 2: Heat = 700; Fil = 4; Vel = 55; Del = 130; Pul = 55; 1-step pulling. 
		NOTE: The units are specific for each puller and glass capillary used here, respectively (see Table of Materials). Other capillaries and pullers can also be used to prepare the glass needles. But the glass needles should not be too thin as they might break when coming in contact with the skull. Capillary: length: 100 mm (4 inches); OD: 1.5 mm; ID: 0.84 mm; filament: Yes</li>
	</ol>
	</li>
</ol>
2. Anesthesia of larvae and preparations for embedding
<ol>
	<li>When starting the experiment for the day, transfer the animals that are needed with a plastic Pasteur pipette to a 90 mm diameter Petri dish, which is filled with either Danieau (for larvae which are still kept in a Petri dish with Danieau) or water from the fish facility (for larvae which are older than 7 dpf and are kept in the fish facility).
	<ol>
		<li>When pipetting fish older than 2 weeks, make sure the opening of the pipette is large enough to avoid injuring the fish when transferring them. Do not use a net because it will physically damage especially the younger larvae.</li>
	</ol>
	</li>
	<li>Add rotifera or artemia nauplii suited for the size of the larvae kept in the Petri dish, to ensure free access to food and maximum health status of the larvae and to reduce stress.</li>
	<li>For embedding, transfer the selected larvae to a 35 mm diameter Petri dish filled with ACSF. Add the necessary volume of d-Tubocurarine to reach a working concentration/effective dose of 10 &#181;M and wait for a few minutes until the larvae are completely immobilized11. 
	NOTE: When the fish grow older or if a faster full anesthesia is needed (under 5 min), it is possible to increase the concentration of d-Tubocurarine (LD50 for mice is 0.13 mg/kg intravenously12). It is also possible to use a different anesthetic, such as &#945;-bungarotoxin (working concentration: 1 mg/mL), which has the same effect as curare and also keeps the brain fully active13. If a fully active brain is not necessary for the subject of interest, Tricaine in a non-lethal dose (0.02%) is also an option to fully anesthetize the larvae. However, Tricaine blocks sodium-channels, thereby impairing brain activity14.</li>
	<li>Prepare the mounting chamber by taking the lid of the 35 mm diameter Petri dish, flip the lid upside down, and place a square glass coverslip (24 x 24 mm) on the bottom of the lid. See Figure 1 (upper part) for a schematic description of these steps. The smoother surface of the glass prevents slipping away of the agarose block, which contains the larvae during the skull opening procedure.</li>
	<li>Aliquot the amount of ACSF needed for the day in an appropriate vial (e.g., 50 mL tube, beaker, Schott bottle, etc.) and oxygenate it with carbogen (5% CO2, 95% O2). If imaging only morphology (e.g., fluorescence patterns) ACSF is still necessary to ensure the integrity of the brain and that cells are not negatively influenced by osmolarity effects but oxygenation of the ACSF is not needed. This step only needs to be performed&#160;when full brain activity is necessary for imaging. 
	NOTE: For optimal oxygen saturation of the medium, add an air stone to the end of the carbogen tube. To guarantee a sufficiently high oxygen level, it is necessary to exchange the ACSF in the imaging chambers with freshly oxygenated ACSF every 20-60 min, depending on the number and age of larvae embedded in the same imaging chamber (e.g., for a single embedded larva&#160;ACSF exchange every hour is sufficient. For six larvae older than 14 dpf embedded in parallel, exchanging ACSF every 20 min is necessary) so plan the necessary amount of oxygen saturated ACSF according to the planned experiment.</li>
</ol>
3. Embedding of the larvae
<ol>
	<li>Transfer the fully anesthetized larvae with a plastic Pasteur pipette to the (in step 2.4) prepared mounting chamber. Then, carefully remove the excess medium to avoid dilution of the LM-agarose. All the following steps should be performed under a stereo microscope with sufficient magnification. 
	NOTE: Tilting the mounting chamber can help to fully remove the medium.</li>
	<li>Proceed immediately to the next step, by adding a sufficiently large LM-agarose drop on top of the larvae (circa 1 mL, depending on the size of the larvae) to protect the animals from drying out and to reduce unnecessary stress.</li>
	<li>Orient the larvae in position before the agarose solidifies. Ensure that the dorsal part of the larvae is directed upward. Also, make sure to embed the larvae as close to the surface of the agarose as possible. 
	NOTE: Depending on the size and number of larvae planned to embed at the same time, it is possible to adjust the agarose concentration. For example, for 1-3 larvae that are 30 dpf old, a concentration of 1.8%-2% LM-agarose is recommended. For 1-4 larvae that are 7 dpf old, it is most practicable to use 2.5% LM-agarose, whereas, for 5-8 larvae, 2% is more suited. If a fully active brain is required, it is recommended to only embed three fish at the same time to reduce the time needed to operate the larvae. In general, it is recommended to use lower concentrations (1.8%-2%) the older the larvae get or the more larvae are planned to be embedded at the same time.</li>
	<li>If images will be recorded using an inverted microscope, trim the agarose block containing the larvae into a small cuboid shape. This is important for transferring the larvae to the imaging chamber later on. If using an upright microscope, such trimming is not necessary, because the mounting chamber can also be used as the imaging chamber. In Figure 1 (upper part), one can find a schematic description of these steps.</li>
</ol>
4. Exposing the brain
NOTE: All the following steps should be performed with greatest care to not unnecessarily injure the larvae. If a fully active brain is required for the experiment, keep in mind that with every second that passes, while the fish is still fully mounted in agarose and has an open skull without oxygenated ACSF, the brain will suffer from a lack of oxygen and also dry out. The effects of oxygen deficiency will become even more dramatic, the older the embedded larvae are. Therefore, it is important to perform the surgery not only within the shortest time possible, but also with maximum precision to not evoke mechanical brain damage with the needle. When trained, steps 4.2-4.4 should not take more than 30 s per fish.
<ol>
	<li>Begin the surgery as soon as the agarose has solidified. First, trim away all of the excess agarose above the brain region of interest to obtain free access to the head and a clear working space. If the dorsal part of the head is already sticking out of the agarose, skip this step.</li>
	<li>Depending on the region of interest, pick a spot to begin with the surgery. Take the glass needle and make a small incision through the skin but without penetrating too deep into the tissue. This will be the starting point for peeling away the overlaying skin. 
	NOTE: For optimal results, never start directly above the region of interest to reduce the risk of damaging important structures. If necessary, it is possible to even start posterior to the hindbrain and from there work forward until the unwanted area of skin is peeled away.</li>
	<li>Continue with very small cuts along the part of skin aiming to remove by barely moving the needle just underneath the surface. Most of the time it is not necessary to move completely around the brain and to cut out a circle-like piece of skin and skull, but rather just make two incisions along the head and then push the skin away to one or the other side. Figure 2 shows a schematic representation of the optimal cutting strategy to obtain free access to the cerebellum. 
	NOTE: This micro-surgery is a delicate procedure and it will most likely need some training to perfectly remove the skin without damaging the underlying brain. It is also recommended to find out the optimal cutting strategy for the brain region of interest and stick with it for the period of the experiment.</li>
	<li>Immediately after removing the skin from all embedded larvae, proceed by pouring (oxygenated) ACSF over the agarose to flood away unwanted skin particles and blood and to keep the brain fully active and protect it from drying out. 
	NOTE: If a healthy brain is needed for the experiment, it is recommended to go for a maximum of three fish at a time.</li>
	<li>If using an upright microscope, start directly with imaging.
	<ol>
		<li>When using an inverted microscope, slide a small spatula underneath the cuboid agarose block (step 3.4).</li>
		<li>Add a small drop of LM-agarose to the bottom of the imaging chamber (e.g., glass-bottom dish) and immediately flip the agarose block containing the larvae with the spatula for 180&#176; and gently push it to the bottom of the imaging chamber, while the liquid agarose drop acts as glue.</li>
		<li>When the agarose has solidified, fill up the imaging chamber with (oxygenated) ACSF, then begin imaging. See Figure 1 (lower part) for a schematic description.</li>
	</ol>
	</li>
	<li>When full brain activity is required for the experiment, always make sure that ACSF in the imaging chamber has a sufficiently high oxygen level. To ensure this, exchange the medium carefully with freshly oxygenated ACSF when possible every 20-60 min (depending on the number and size of the fish, size and surface of the imaging chamber, and imaging duration).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62166/62166fig01.jpg" /> 
Figure 1:&#160;Schematic procedure for the preparation of open skull zebrafish for in vivo imaging in a stepwise manner. The working instructions for the different steps can be found in the graphic itself. Graphic designed by Florian Hetsch and adapted by Paul Schramm. <a href="https://www.jove.com/files/ftp_upload/62166/62166fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62166/62166fig02.jpg" /> 
Figure 2:&#160;Detailed schematic representation of the micro-surgery performed to remove pieces of skin and skull above the brain region of interest. The red circle marks the spot where the first cut needs to be made. The red-dotted line delineates the optimal path to cut along with the needle to obtain free access to the cerebellum without damaging it. The green arrow marks the direction in which the excessive skin and skull pieces can easily be pushed away. Make sure to never penetrate into the brain tissue during the whole procedure. After successfully peeling away the skin, the brain region of interest (here, cerebellum) will be freely accessible for any kind of high-resolution in vivo imaging. <a href="https://www.jove.com/files/ftp_upload/62166/62166fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

The presented method provides an alternative approach to brain isolation or the treatment of zebrafish larvae with pharmaceuticals inhibiting pigmentation for recording high resolution images of neurons in their in vivo environment. The quality of images recorded with this method is comparable to images from explanted brains, yet under natural conditions.
Furthermore, a loss in intensity of fluorescence is avoided, because there is no need for treatment with fixatives<sup class="xref"...

Over the recent decades, the zebrafish (Danio rerio) has evolved as one of the most popular vertebrate model organisms for embryonic and larval developmental studies. The large fecundity of zebrafish females coupled with the rapid ex utero development of the embryo and its transparency during early embryonic developmental stages are just a few key factors that make zebrafish a powerful model organism to adress developmental questions1. Advances in molecular genetic technologies combined with high resolution in vivo imaging studies allowed for addressing cell biological mechanisms underlying developmental processes2. In particular, in the field of neuronal differentiation, physiology, connectivity, and function, zebrafish has shed light on the interplay of molecular dynamics, brain functions and organismic behavior in unprecedented detail.
Yet, most of these studies are restricted to embryonic and early larval stages during the first week of development as transparency of the nervous system tissue is progressively lost. At these stages, brain tissue is prevented from access by high resolution microscopy approaches becoming shielded by skull differentiation and pigmentation3.
Therefore, key questions of neuronal differentiation, maturation, and plasticity such as the refinement of neuronal connectivity or synaptic scaling are difficult to study. These cellular processes are important in order to define cellular mechanisms driving, for example, social behavior, decision making, or motivation-based behavior, areas to which zebrafish research on several weeks&#39; old larvae has recently contributed key findings based on behavioral studies4.
Pharmacological approaches to inhibit pigmentation in zebrafish larvae for several weeks are barely feasible or may even cause detrimental effects5,6,7,8. Double or triple mutant strains with specific pigmentation defects, such as casper9 or crystal10, have become tremendously valuable tools, but are laborious in breeding, provide few offspring, and pose the danger of accumulating genetic malformations due to excessive inbreeding.
Here, a minimal invasive procedure as an alternative is provided that is applicable to any zebrafish strain. This procedure was adapted from electrophysiological studies to record neuronal activity in living and awake zebrafish larvae. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, because they are not tightly interwoven with the brain vasculature. This allows for exposing brain tissue containing neurons and axonal tracts without damage and for recording neuronal morphology, including synaptic structures and their molecular contents, which in turn include the observation of physiological changes such as Ca2+ transients or intracellular transport events for up to several hours. Moreover, beyond descriptive characterizations, the direct access to brain tissue enables interrogation of mature neuronal functions by means of neuropharmacological substance administration and optogenetic approaches. Therefore, true structure function relationships can be revealed in the juvenile zebrafish brain using this brain exposure strategy.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Calcium chloride</td>
			<td>Roth</td>
			<td>A119.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Laser scanning microscope</td>
			<td>Leica</td>
			<td>TCS SP8</td>
			<td></td>
		</tr>
		<tr>
			<td>d-Glucose</td>
			<td>Sigma</td>
			<td>G8270-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>d-Tubocurare</td>
			<td>Sigma-Aldrich</td>
			<td>T2379-100MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Capillary type 1</td>
			<td>WPI</td>
			<td>1B150F-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Capillary type 2</td>
			<td>Harvard Apparatus</td>
			<td>GC100F-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Coverslip</td>
			<td>deltalab</td>
			<td>D102424</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Roth</td>
			<td>9105.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoechst 33342</td>
			<td>Invitrogen (Thermo Fischer)</td>
			<td>H3570</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging chamber</td>
			<td>Ibidi</td>
			<td>81156</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Normapur</td>
			<td>26764298</td>
			<td></td>
		</tr>
		<tr>
			<td>LM-Agarose</td>
			<td>Condalab</td>
			<td>8050.55</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride (Hexahydrate)</td>
			<td>Roth</td>
			<td>A537.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Camera</td>
			<td>Leica</td>
			<td>DFC9000 GTC</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle-Puller type 1</td>
			<td>NARISHIGE</td>
			<td>Model PC-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle-Puller type 2</td>
			<td>Sutter Instruments</td>
			<td>Model P-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur-Pipettes 3ml</td>
			<td>A.Hartenstein</td>
			<td>20170718</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Roth</td>
			<td>P029.2</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Normapur</td>
			<td>28244262</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricain</td>
			<td>Sigma-Aldrich</td>
			<td>E10521-50G</td>
			<td></td>
		</tr>
		<tr>
			<td>Waterbath</td>
			<td>Phoenix Instrument</td>
			<td>WB-12</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm petri dish</td>
			<td>Sarstedt</td>
			<td>833900</td>
			<td></td>
		</tr>
		<tr>
			<td>90 mm petri dish</td>
			<td>Sarstedt</td>
			<td>821473001</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo imaging of fully active brain tissue in awake zebrafish larvae and juveniles by skull and skin removal

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at cellular and subcellular resolution. Advances in molecular and imaging technologies have allowed us to gain numerous detailed insights into cellular and molecular mechanisms of brain development in the transparent zebrafish embryo. Recently, processes of refinement of neuronal connectivity that occur at later larval stages several weeks after fertilization, which are for example control of social behavior, decision making or motivation-driven behavior, have moved into focus of research. At these stages, pigmentation of the zebrafish skin interferes with light penetration into brain tissue, and solutions for embryonic stages, e.g., pharmacological inhibition of pigmentation, are not feasible anymore.
Therefore, a minimally invasive surgical solution for microscopy access to the brain of awake zebrafish is provided that is derived from electrophysiological&#160;approaches. In teleosts, skin and soft skull cartilage can be carefully removed by micro-peeling these layers, exposing underlying neurons and axonal tracts without damage. This allows for recording neuronal morphology, including synaptic structures and their molecular contents, and the observation of physiological changes such as Ca2+ transients or intracellular transport events. In addition, interrogation of these processes by means of pharmacological inhibition or optogenetic manipulation is feasible. This brain exposure approach provides information about structural and physiological changes in neurons as well as the correlation and interdependence of these events in live brain tissue in the range of minutes or hours. The technique is suitable for in vivo brain imaging of zebrafish larvae up to 30 days post fertilization, the latest developmental stage tested so far. It, thus, provides access to such important questions as synaptic refinement and scaling, axonal and dendritic transport, synaptic targeting of cytoskeletal cargo or local activity-dependent expression. Therefore, a broad use for this mounting and imaging approach can be anticipated.

Figure 3A,C show a 14 dpf larva of the transgenic line Tg[-7.5Ca8:GFP]bz12[15] with the skull still intact. The pigment cells in the overlaying skin are distributed all over the head and are interfering with the fluorescence signal in the region of interest (here, cerebellum). With the larva in this condition, it is not possible to obtain high resolution images of the brain. F...

Watch this Scientific Journal Video about In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal at JoVE.com

In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Understanding the ephemeral changes that occur during brain development and maturation requires detailed high-resolution imaging in space and time at ...

in-vivo-imaging-fully-active-brain-tissue-awake-zebrafish-larvae

Research

JoVE Journal

Neuroscience

4.6K Views.  Technische Universität Braunschweig. Here we present a method to image the zebrafish embryonic brain in vivo upto larval and juvenile stages. This microinvasive procedure, adapted from electrophysiological approaches, provides access to cellular and subcellular details of mature neuron and can be combined with optogenetics and neuropharmacological studies for characterizing brain function and drug intervention.

Video: In vivo Imaging of Fully Active Brain Tissue in Awake Zebrafish Larvae and Juveniles by Skull and Skin Removal

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

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

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

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

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

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

Exposure of the Pig CNS for Histological Analysis: A Manual for Decapitation, Skull Opening, and Brain Removal

Pigs have become increasingly popular in large-animal translational neuroscience research as an economically and ethically feasible substitute to non-human primates. The large brain size of the pig allows the use of conventional clinical brain imagers and the direct use and testing of neurosurgical procedures and equipment from the human clinic. Further macroscopic and histological analysis, however, requires postmortem exposure of the pig central nervous system (CNS) and subsequent brain removal. This is not an easy task, as the pig CNS is encapsulated by a thick, bony skull and spinal column. The goal of this paper and instructional video is to describe how to expose and remove the postmortem pig brain and the pituitary gland in an intact state, suitable for subsequent macroscopic and histological analysis.

The goal of this paper and instructional video is to describe how to expose and remove the postmortem pig brain and pituitary gland in an intact state, suitable for subsequent macroscopic and histological analysis.

Pigs have become increasingly popular in large-animal translational neuroscience research as an economically and ethically feasible substitute to ...

Labeling and Imaging of Amyloid Plaques in Brain Tissue Using the Natural Polyphenol Curcumin

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). Therefore, detection of the presence of A&#946; in AD brain tissue is a valuable tool for developing new treatments to prevent the progression of AD. Several classical amyloid binding dyes, fluorochrome, imaging probes, and A&#946;-specific antibodies have been used to detect A&#946; histochemically in AD brain tissue. Use of these compounds for A&#946; detection is costly and time consuming. However, because of its intense fluorescent activity, high-affinity, and specificity for A&#946;, as well as structural similarities with traditional amyloid binding dyes, curcumin (Cur) is a promising candidate for labeling and imaging of A&#946; plaques in postmortem brain tissue. It is a natural polyphenol from the herb Curcuma longa. In the present study, Cur was used to histochemically label A&#946; plaques from both a genetic mouse model of 5x familial Alzheimer&#39;s disease (5xFAD) and from human AD tissue within a minute. The labeling capability of Cur was compared to conventional amyloid binding dyes, such as thioflavin-S (Thio-S), Congo red (CR), and Fluoro-jade C (FJC), as well as A&#946;-specific antibodies (6E10 and A11). We observed that Cur is the most inexpensive and quickest way to label and image A&#946; plaques when compared to these conventional dyes and is comparable to A&#946;-specific antibodies. In addition, Cur binds with most A&#946; species, such as oligomers and fibrils. Therefore, Cur could be used as the most cost-effective, simple, and quick fluorochrome detection agent for A&#946; plaques.

Curcumin is an ideal fluorophore for labeling and imaging of amyloid beta protein plaques in brain tissue due to its preferential binding to amyloid beta protein as well as its structural similarities with other traditional amyloid binding dyes. It can be used to label and image amyloid beta protein plaques more efficiently and inexpensively than traditional methods.

Deposition of amyloid beta protein (A&#946;) in extra- and intracellular spaces is one of the hallmark pathologies of Alzheimer&#39;s disease (AD). ...

A Hydrophobic Tissue Clearing Method for Rat Brain Tissue

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered tissue samples. Traditional immunolabeling procedures require cutting the sample into thin sections, which restricts the ability to label and examine intact structures. However, if brain tissue can remain intact during processing, structures and circuits can remain intact for the analysis. Previously established clearing methods take significant time to completely clear the tissue, and the harsh chemicals can often damage sensitive antibodies. The iDISCO method quickly and completely clears tissue, is compatible with many antibodies, and requires no special lab equipment. This technique was initially validated for the use in mice tissue, but the current protocol adapts this method to image hemispheres of control and transgenic rat brains. In addition to this, the present protocol also makes several adjustments to preexisting protocol to provide clearer images with less background staining. Antibodies for Iba-1 and tyrosine hydroxylase were validated in the HIV-1 transgenic rat and in F344/N control rats using the present hydrophobic tissue clearing method. The brain is an interwoven network, where structures work together more often than separately of one another. Analyzing the brain as a whole system as opposed to a combination of individual pieces is the greatest benefit of this whole brain clearing method.

Here we present a hydrophobic tissue clearing method that allows for the viewing of target molecules as part of intact brain structures. This technique has now been validated for F344/N control and HIV-1 transgenic rats of both sexes.

Hydrophobic tissue clearing methods are easily adjustable, fast, and low-cost procedures that allows for the study of a molecule of interest in unaltered ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the ...

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their activity in vivo in behaving mice. This paper describes a method for the implantation of a chronic cranial window to allow for the longitudinal imaging of brain cells in awake, head-restrained mice. In combination with genetic strategies and viral injections, it is possible to label specific cells and regions of interest with structural or physiological markers. This protocol demonstrates how to combine viral injections to label neurons in the vicinity of GCaMP6-expressing astrocytes in the cortex for simultaneous imaging of both cells through a cranial window. Multiphoton imaging of the same cells can be performed for days, weeks, or months in awake, behaving animals. This approach provides researchers with a method for viewing cellular dynamics in real time and can be applied to answer a number of questions in neuroscience.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their ...