This work was supported by the National Institutes of Health (NIH) (R01- NS115947 awarded to F. Hoerndli). We would also like to thank Dr. Attila Stetak for the pAS1 plasmid.

1. Creating transgenic strains
<ol>
	<li>Using a cloning method of choice8,9, clone expression vectors to contain the Pflp-18 or Prig-3 promoter (for AVA-specific signal in the ventral nerve cord), followed by the Ca2+ indicator of choice, and then a 3&#39; UTR (see the discussion for more information)10. A list of plasmids and their sources can be found in Supplemental Table 1.</li>
	<li>Follow the established protocol for the creation of transgenic strains via the microinjection of a DNA mix (&#60;100 ng/&#956;L; see Supplemental Table 1 for the concentrations of each plasmid) containing the Ca2+ indicator transgene (~20 ng/&#956;L) and the rescue DNA lin-15+ (selection marker) into the gonad of a 1 day old adult C. elegans10 (genotype: lin-15(n765ts); phenotype: multi-vulva)10,11. 
	NOTE. Maintain lin-15(n765ts) parents at 15 &#176;C to suppress temperature-sensitive mutation.</li>
	<li>Maintain the injected parents at 20 &#176;C until the F1 progeny reach adulthood to allow for transgenic selection using the absence of the multi-vulva phenotype.</li>
	<li>Clonally passage13 adult F1 progeny that do not have the multi-vulva phenotype as this indicates the expression of the extrachromosomal array containing the Ca2+ indicator11.</li>
	<li>Assess the expression of the Ca2+ indicator in the F2 or F3 progeny that do not have the multi-vulva phenotype by mounting and imaging 1 day old adults as described later in section 3. 
	NOTE. A relatively high expression of the indicator is required for the rapid acquisition of images as described here (see the discussion for more detail).</li>
	<li>Perform experiments on subsequent generations of the transgenic strains with an optimal expression of the Ca2+ indicator (see the discussion for more detail), maintaining the strains under standard conditions (20 &#176;C on NGM plates)12.</li>
</ol>
2. Optical setup
<ol>
	<li>Use a microscope capable of long-term time-lapse imaging. 
	NOTE. The representative data were acquired using an inverted spinning disk confocal microscope equipped with 488 nm and 561 nm excitation lasers.</li>
	<li>Use a low-magnification objective (i.e., 10x/0.40) for the crude localization of the worm.</li>
	<li>To achieve subcellular resolution, switch to and localize the neurons using a high-magnification objective. 
	NOTE. A 100x/1.40 NA oil objective was used to acquire the representative data in this study.</li>
	<li>Acquire the images using an ultra-sensitive camera capable of rapid image acquisition (&#62;50 fps).</li>
	<li>Use a standard emission filter for the Ca2+ indicator of choice (i.e., a 525 nm &#177; 50 nm emission filter for GCaMP6f and mitoGCaMP).</li>
	<li>Add a Z-drift corrector (ZDC) for the acquisition of image streams longer than 10 s, as the desiccation of the agar pad during the imaging session causes the neurite to drift out of focus within several seconds. 
	NOTE. If a microscopy set-up does not allow for ZDC, or if long (&#62;20 min) imaging periods are desired, then apply a border of silicon grease or melted petroleum jelly around the agar pad to slow the desiccation.</li>
</ol>
3. Preparation of worms for imaging
<ol>
	<li>Preparing the agar pads
	<ol>
		<li>Make 3 mL of 10% agar by dissolving molecular grade agar in M912 in a 13 mm x 100 mm glass culture tube and microwaving for several seconds (see Table of Materials). 
		NOTE. Molten agar can be kept on a heat block for up to 1 h or can be made fresh each time agar pads are needed.</li>
		<li>Place a microscope slide between two additional slides that each have two layers of laboratory tape (see Figure 1A-i).</li>
		<li>Cut the tip of a 1,000 &#956;L pipette tip (without a filter), and use it to pipette a small drop of agar onto the center coverslip (Figure 1A-i).</li>
		<li>Flatten the agar by pressing another slide down on top of the agar (Figure 1A-ii).</li>
		<li>After cooling, cut the agar into a small disc using the opening of a 13 mm x 100 mm glass culture&#160;tube (Figure 1A-iii), and then remove the surrounding agar.</li>
	</ol>
	</li>
	<li>Preparing the worm-rolling solution
	<ol>
		<li>Dissolve muscimol powder in M9 to create a 30 mM stock. Separate into 50 &#956;L aliquots, and store at 4 &#176;C.</li>
		<li>Thaw a new aliquot of 30 mM muscimol every 3-5 days (and store at 20 &#176;C while in use).</li>
		<li>Dilute a 30 mM muscimol stock in a 1:1 ratio with polystyrene beads to make the rolling solution.</li>
	</ol>
	</li>
	<li>Positioning a worm for imaging
	<ol>
		<li>Place 1.6 &#956;L of the rolling solution onto the center of the agar pad. 
		NOTE: The amount of liquid should be adjusted for imaging younger (less) or older animals (more).</li>
		<li>Using the preferred worm pick (i.e., a glass or platinum wire pick)13, transfer a worm of the desired age without the multi-vulva phenotype11 into the rolling solution on the agar pad (Figure 1A-iv). 
		NOTE. The representative data are from 1 day old hermaphrodites (GCaMP6f strain = FJH185; mitoGCaMP strain; FJH597; see Supplemental Table 1), which can be identified by the presence of only one row of eggs. See the discussion for more detail on working with worms of different ages.</li>
		<li>Wait for ~5 min for the muscimol to reduce the worm movement, and then drop a 22 mm x 22 mm coverslip on top of the agar pad, physically restricting the worm movement.</li>
		<li>For imaging neurites in the ventral nerve cord, roll the worm into the orientation shown in Figure 1B,C by lightly sliding the coverslip. 
		NOTE. To visualize the ventral nerve cord, position the worm with the head upward, and roll the worm until the intestine is on the right side of the proximal portion and the left for the distal portion of the worm (from the viewer&#39;s perspective). Invert this orientation (Figure 1C) for imaging neurites in the dorsal nerve cord.</li>
	</ol>
	</li>
	<li>Mounting the worm on a microscope
	<ol>
		<li>Place a drop of immersion oil onto the coverslip before mounting it onto the microscope stage.</li>
		<li>Find the worm using a low-magnification objective (10x) objective in brightfield.</li>
		<li>Switch to the 100x objective, and adjust the focus.</li>
		<li>Locate the AVA neurite using the illumination of the GCaMP or mitoGCaMP with the 488 nm imaging laser. 
		NOTE: Use small adjustments to prevent squishing the worm or pulling off the coverslip.</li>
	</ol>
	</li>
</ol>
4. Acquisition of high-resolution image streams
<ol>
	<li>In vivo GCaMP imaging
	<ol>
		<li>Set the exposure time to 20 ms.</li>
		<li>Adjust the imaging laser and acquisition settings until the basal GCaMP fluorescence is in the mid-range of the camera&#39;s dynamic range14. Refer to the discussion for suggestions on optimizing the imaging parameters using other microscopy setups. 
		&#8203;NOTE. For the optical setup used in this study, set the 488 nm imaging laser to the following output settings: laser power = 50% and attenuation = 1. Modify the additional acquisition settings as follows: EM gain = 200 and pre-amplifier gain = 1.</li>
		<li>After the AVA neurite is located using the GCaMP fluorescence at 100x magnification, set the ZDC (see Table of Materials) to a range of &#177;30 &#956;m, and initiate continuous autofocusing. Manually correct the focal plane as needed before imaging.</li>
		<li>Acquire a stream of images at 100x magnification. Durations as long as 10 min can be achieved with the setup used in this study.</li>
	</ol>
	</li>
	<li>In vivo mitoGCaMP imaging
	<ol>
		<li>Follow steps 4.1.1-4.1.2 as needed to acquire the basal mitoGCaMP fluorescence in the mid-range for the camera&#39;s dynamic range14. 
		NOTE. For the optical set-up used in this study, set the 488 nm imaging laser to the following output settings: laser power = 20% and attenuation = 10. Modify the acquisition settings to the following: EM gain = 100 and pre-amplifier gain = 2.</li>
		<li>After the mitochondria in the AVA neurite are located using the mitoGCaMP fluorescence, set the ZDC as described above in step 3.3.</li>
		<li>Acquire a stream of images at 100x magnification.</li>
	</ol>
	</li>
</ol>
5. Image stream analysis
<ol>
	<li>Open the image stream in an appropriate image software for analyzing the pixel values in an image series (see Table of Materials).</li>
	<li>Using the Polygon Selection tool, define the subcellular region of interest (ROI) containing GCaMP or mitoGCaMP.</li>
	<li>Click on Analyze &#62; Tools &#62; ROI Manager. In the ROI Manager, click Add and then More &#62; Multi Measure. In the Multi Measure window, click OK to automatically collect the average, minimum, and maximum pixel intensities of the ROI defined above for each frame of the image stream for further analysis. 
	NOTE. It is recommended that the average pixel intensity (Favg) is normalized to the minimum intensity (Fmin) for each ROI using the ratio Favg/Fmin to account for fluorescence differences due to positioning in the z-plane. Discard the image streams with worm movement during imaging since this would also affect the fluorescence measurements.</li>
</ol>

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The authors declare no competing interests.

The first consideration when implementing the method described involves the selection of a Ca2+ indicator with ideal characteristics for the given research question. Cytoplasmic Ca2+ indicators typically have a high affinity for Ca2+,&#160;and the sensitivity of these indicators to Ca2+ is inversely related to the kinetics (on/off rate)16,17. This means that either Ca2+ sensitivity or kinetics will need to be...

Calcium ions (Ca2+) are highly versatile carriers of information in many cell types. In neurons, Ca2+ is responsible for the activity-dependent release of neurotransmitters, the regulation of cytoskeletal motility, the fine-tuning of metabolic processes, as well as many other mechanisms required for proper neuronal maintenance and function1,2. To ensure effective intracellular signaling, neurons must maintain low basal Ca2+ levels in their cytoplasm3. This is accomplished by cooperative Ca2+ handling mechanisms, including the uptake of Ca2+ into organelles such as the endoplasmic reticulum (ER) and mitochondria. These processes, in addition to the arrangement of Ca2+-permeable ion channels in the plasma membrane, result in heterogeneous levels of cytoplasmic Ca2+ throughout the neuron.
Ca2+ heterogeneity during rest and neuronal activation allows for the diverse, location-specific regulation of Ca2+-dependent mechanisms1. One example of the concentration-specific effects of Ca2+ is the release of Ca2+ from the ER through inositol 1,4,5-trisphosphate (InsP3) receptors. Low Ca2+ levels in combination with InsP3 are required for the opening of the receptor&#39;s calcium-permeable pore. Alternatively, high Ca2+ levels both directly and indirectly inhibit the receptor4. The importance of Ca2+ homeostasis for proper neuronal function is supported by evidence suggesting that disrupted intracellular Ca2+ handling and signaling is an early step in the pathogenesis of neurodegenerative disorders and natural aging5,6. Specifically, abnormal Ca2+ uptake and release by the ER and mitochondria are linked to the onset of neuronal dysfunction in Alzheimer&#39;s disease, Parkinson&#39;s disease, and Huntington&#39;s disease6,7.
The study of Ca2+ dyshomeostasis during natural aging or neurodegeneration requires the longitudinal observation of Ca2+ levels with subcellular resolution in a living, intact organism in which the native cellular architecture (i.e., the arrangement of synapses and distribution of ion channels) is maintained. To this end, this protocol provides guidance on the use of two readily available, genetically encoded Ca2+ sensors for recording Ca2+ dynamics in vivo with high spatial and temporal resolution. The representative results acquired using&#160;the described method in C. elegans demonstrate how the expression of Ca2+ indicators in the cytoplasm or mitochondrial matrix of single neurons can allow for the rapid acquisition of fluorescent images (up to 50 Hz) that illustrate the Ca2+ dynamics with the additional capability of discerning the Ca2+ levels within single spine-like structures and individual mitochondria.

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			<td>Beam homogenizer</td>
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			<td>Borealis upgrade to CSU-X1</td>
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			<td>CleanBench laboratory table&#160;</td>
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			<td>ILE laser combiner&#160;</td>
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			<td>4 laser lines&#160;</td>
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			<td>ILE solid state 488 nm laser</td>
			<td>Andor Technologies&#160;</td>
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			<td>50 mW</td>
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			<td>ImageJ</td>
			<td>National Institutes of Health</td>
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			<td>IX83 Spinning disk confocal microscope&#160;</td>
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			<td>With Yokogawa CSU-X1 spinning disc</td>
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			<td>iXon Ultra EMCCD camera&#160;</td>
			<td>Andor Technologies&#160;</td>
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			<td>pJM23</td>
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			<td>Norlake Scientific</td>
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			<td>Set to 15 &#176;C</td>
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			<td>Stage controller</td>
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			<td>Standard microscope slide</td>
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			<td>Touch panel controller</td>
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			<td>Z-drift corrector&#160;</td>
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subcellular imaging of neuronal calcium handling in vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a crucial component of intracellular signaling. The visualization of subcellular Ca2+ dynamics in vivo,&#160;where neurons can be studied in their native, intact circuitry, has proven technically challenging in complex nervous systems. The transparency and relatively simple nervous system of the nematode Caenorhabditis elegans enable the cell-specific expression and in vivo visualization of fluorescent tags and indicators. Among these are fluorescent indicators that have been modified for use in the cytoplasm as well as various subcellular compartments, such as the mitochondria. This protocol enables non-ratiometric Ca2+ imaging in vivo with a subcellular resolution that permits the analysis of Ca2+ dynamics down to the level of individual dendritic spines and mitochondria. Here, two available genetically encoded indicators with different Ca2+ affinities are used to demonstrate the use of this protocol for measuring relative Ca2+ levels within the cytoplasm or mitochondrial matrix in a single pair of excitatory interneurons (AVA). Together with the genetic manipulations and longitudinal observations possible in C. elegans, this imaging protocol may be useful for answering questions regarding how Ca2+ handling regulates neuronal function and plasticity.

These two protocols enable the rapid acquisition of differential Ca2+ levels within the subcellular regions and organelles of individual neurites in vivo with high spatial resolution. The first protocol allows for the measurement of cytoplasmic Ca2+ with high temporal and spatial resolution. This is demonstrated here using the cell-specific expression of GCaMP6f in the glutamatergic AVA command interneurons15, whose neurites run the entire length of the ventral nerve...

Watch this Scientific Journal Video about Subcellular Imaging of Neuronal Calcium Handling In Vivo at JoVE.com

Subcellular Imaging of Neuronal Calcium Handling In Vivo

The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a ...

subcellular-imaging-of-neuronal-calcium-handling-in-vivo

Research

JoVE Journal

Neuroscience

1.1K Views.  Colorado State University College of Veterinary Medicine and Biomedical Sciences. The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Video: Subcellular Imaging of Neuronal Calcium Handling In Vivo

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new method of Ca2+-imaging using the bioluminescent reporter GFP-aequorin (GA) has been developed. This new technique relies on the fusion of the GFP and aequorin genes, producing a molecule capable of binding calcium and&#160;&#8212; with the addition of its cofactor coelenterazine &#8212; emitting bright light that can be monitored through a photon collector. Transgenic lines carrying the GFP-aequorin gene have been generated for both mice and Drosophila. In Drosophila, the GFP-aequorin gene has been placed under the control of the GAL4/UAS binary expression system allowing for targeted expression and imaging within the brain. This method has subsequently been shown to be capable of detecting both inward Ca2+-transients and Ca2+-released from inner stores. Most importantly it allows for a greater duration in continuous recording, imaging at greater depths within the brain, and recording at high temporal resolutions (up to 8.3 msec). Here we present the basic method for using bioluminescent imaging to record and analyze Ca2+-activity within the mushroom bodies, a structure central to learning and memory in the fly brain.

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Here we present a novel Ca2+-imaging approach using a bioluminescent reporter. This approach uses a fused construct GFP-aequorin which binds to Ca2+ and emits light, eliminating the need for light excitation. Significantly this method permits long continuous imaging, access to deep brain structures and high temporal resolution.

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new ...

Prolonged Incubation of Acute Neuronal Tissue for Electrophysiology and Calcium-imaging

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the experimental time, and increases the number of animals that are utilized per study. This limitation specifically impacts protocols such as calcium imaging that require prolonged pre-incubation with bath-applied dyes. Exponential bacterial growth within 3 - 4 h after slicing is tightly correlated with a decrease in tissue health. This study describes a method for&#160;limiting the proliferation of bacteria in acute preparations to maintain viable neuronal tissue for prolonged periods of time (&#62;24 h) without the need for antibiotics, sterile procedures, or tissue culture media containing growth factors. By cycling the extracellular fluid through UV irradiation and keeping the tissue in a custom holding chamber at 15 - 16 &#176;C, the tissue shows no difference in electrophysiological properties, or calcium signaling through intracellular calcium dyes at &#62;24 h postdissection. These methods will not only extend experimental time for those using acute neuronal tissue, but will reduce the number of animals required to complete experimental goals, and will set a gold standard for acute neuronal tissue incubation.

Once removed from the body, neuronal tissue is greatly affected by environmental conditions, leading to eventual degradation of the tissue after 6 - 8 h. Using a unique incubation method, which closely monitors and regulates the extracellular environment of the tissue, tissue viability can be significantly extended for &#62;24 h.

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

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

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

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

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

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.

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...

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 ...

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to correlate this with local hemodynamics. This protocol uses transgenic mice that have been prepared with a chronic cranial window and express the genetically encoded calcium indicator, RCaMP1.07, under the &#945;-smooth muscle actin promoter to specifically label mural cells, such as vascular smooth muscle cells and ensheathing pericytes. Steps are outlined on how to prepare a tail vein catheter for intravenous injection of fluorescent dyes to trace blood flow, as well as how to measure brain pericyte calcium and local blood vessel hemodynamics (diameter, red blood cell velocity, etc.) by two photon microscopy in vivo through the cranial window in ketamine/xylazine anesthetized mice. Finally, details are provided for the analysis of calcium fluctuations and blood flow movies via the image processing algorithms developed by Barrett et al. 2018, with an emphasis on how these processes can be adapted to other cellular imaging data.

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

This protocol presents steps to acquire and analyze fluorescent calcium images from brain ensheathing pericytes and blood flow data from nearby blood vessels in anesthetized mice. These techniques are useful for studies of mural cell physiology and can be adapted to investigate calcium transients in any cell type.

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to ...