This work was supported by the National Institute of Health National Institute of Neurological Disorders and Stroke (NINDS) grants R01NS069726 and R01NS094539 to SD. We thank Erica DeMers for the audio recording.

Procedures involving animals have been approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Missouri-Columbia.
1. Construction of DNA plasmids
NOTE: For in vitro and in vivo imaging, DNA plasmids encoding GCaMP5G/6s with astrocyte- and neuron-specific promoters and mitochondrial targeting sequences are constructed.
<ol>
	<li>Insert mitochondrial matrix (MM)-targeting sequence (mito-) ATGT CCGTCCTGAC GCCGCTGCTG CTGCGGGGCT TGACAGGCTC GGCCCGGCGG CTCCCAGTGC CGCGCGCCAA GATCCATTCG TTG17 into the cloning sites EcoRI and BamHI in the backbone of adeno-associated virus (AAV) plasmid pZac2.1 to obtain plasmids containing astrocytic gfaABC1D promoter or neuronal CaMKII promoter18,19,20.</li>
	<li>Subclone GCaMP5G/6s into the cloning sites BamH I and Not 1 in the above plasmids to obtain new plasmids pZac-gfaABC1D-mito-GCaMP5G/6s and pZac-CaMKII-mito-GCaMP5G/6s that target transgene expression in mitochondria in astrocytes and neurons20,21 (Figure 1A).</li>
	<li>Prepare pZac-gfaABC1D-mito-GCaMP5G and pZac-CaMKII-mito-GCaMP6s DNA plasmids for transfection for in vitro study (Section 2). Produce AAV vectors with serotype 5 for astrocytes and serotype 9 for neurons for in vivo study18 (Section 3).</li>
</ol>
2. In vitro mitochondrial Ca2+ imaging in astrocytes and neurons
<ol>
	<li>Prepare primary astrocytes from the cortex of P1 neonatal mice and primary neurons from the cortex of E15-16 embryos18,22,23,24, and culture them on 12 mm diameter glass coverslips in 24-well plates using Dulbecco&#39;s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum (FBS), and neuronal basal medium (NBM) containing 2% B27, respectively.</li>
	<li>Transfect mature astrocytes and neurons with pZac-gfaABC1D-GCaMP6s and pZac-CaMKII-GCaMP5G plasmids using lipid based transfection reagent to express GCaMP6s in the mitochondria of astrocytes and GCaMP5G in the mitochondria of neurons18,20,21. Transfect cells in each well with 0.5 &#181;g of DNA, and change the medium 6 h later. 
	NOTE: The astrocytes and neurons are ready for imaging 1-2 days after transfection.</li>
	<li>Perform in vitro mitochondrial Ca2+ imaging 1-2 days after transfection.
	<ol>
		<li>Transfer the glass coverslips cultured with astrocytes or neurons to the PH-1 perfusion chamber under an epifluorescence or two photon microscope.</li>
		<li>Stimulate astrocytic mitochondrial Ca2+ uptake with 100 &#956;M ATP in ACSF, or stimulate neuronal mitochondria with 100 &#956;M glutamate/10 &#956;M glycine20,25 (Figure 2 and Figure 3). 
		NOTE: Solution changes from ACSF to ATP- and glutamate/glycine-containing ACSF are controlled by an ALA-VM8 perfusion system21. The speed of the solution change is controlled at 1-2 mL/min by adjusting a valve.</li>
	</ol>
	</li>
</ol>
3. In vivo mitochondrial Ca2+ imaging in astrocytes and neurons
<ol>
	<li>AAV preparations.
	<ol>
		<li>Prepare the following recombinant adeno-associated virus (rAAV) vectors using the DNA plasmids prepared in section 1: rAAV2/5-gfaABC1D-mito-GCaMP5G and rAAV2/9-CaMKII-mito-GCaMP6s vectors. 
		NOTE: In this experiment, rAAV vectors of serotype 5 were prepared to express GCaMP5G in mitochondria in astrocytes and rAAV vectors of serotype 9 were prepared to express GCaMP6s in mitochondria in neurons.</li>
	</ol>
	</li>
	<li>Stereotaxic AAV injection.
	<ol>
		<li>Anesthetize the mouse with 3% isoflurane. 
		NOTE: Later during the surgery, the isoflurane levels are reduced to 2%.</li>
		<li>After the mouse reaches a surgical level of anesthesia, as determined by tail and toe pinch, shave the hair over the surgery site, motor or somatosensory cortex, with a hair trimmer.</li>
		<li>Position the mouse on the mouse stereotaxic device and fix the head with ear bars. Apply ophthalmic ointment to the eyes to protect them during the surgery. Use a heating pad to keep the body temperature of the mouse at 37 &#176;C throughout the surgery. 
		NOTE: Perform surgery using aseptic procedures. All surgical tools need to be sterilized either by autoclaving or a hot bead sterilizer.</li>
		<li>After the mouse is mounted on the stereotaxic device, sterilize the scalp with alternating iodine based scrub and 70% ethanol three times. Make an incision in the midline of the scalp to expose the injection site.</li>
		<li>Cut open the skin in the bregma-lamda axis and a create a ~1 mm diameter burr hole with a high speed drill at the intended injection location of the motor or somatosensory cortex.</li>
		<li>Use a 33 G Hamilton syringe containing adeno-associated virus (rAAV2/5-gfaABC1D-mito-GCaMP5G vectors [1 x 1011 GC] or rAAV2/9-CaMKII-mito-GCaMP6s vectors [1 x 1011 GC])&#160;to inject up to 1 &#181;L of virus at the target area. 
		NOTE: For example, for cortical viral delivery, inject the virus solution at two depths in multiple steps. First, insert the needle to a depth of 1 mm from the dura&#160;and allow 5 min for the brain to recover. Then, move the needle up to ~700 &#181;m depth and inject 500 nL of the virus solution at an injection speed of 10 nL/s using a hamilton syringe controlled by a microsyringe pump controller. After the injection is completed, wait for 5 min to allow the virus to diffuse into the brain. Then, move the needle up to the second injection location at a depth of 300 &#181;m. Here, inject an additional 500 nL of the virus solution. Wait for 10 min to allow the virus to diffuse into the brain.</li>
		<li>Close the scalp and the skin using a tissue adhesive. Let the mice recover on the heating pad. Send mice back to the animal facility after recovery.</li>
	</ol>
	</li>
	<li>Cranial-window intstallation and in vivo&#160;two-photon (2-P) imaging of mitochondrial Ca2+ signals. 
	NOTE: Cranial window implantation is done 3 weeks post AAV injection over the motor or somatosensory cortex26,27,28,29. Carprofen (10 mg/kg) is injected&#160;subcutaneously to provide relief from potential pain before surgery. The cranial window surgical procedures are identical to the AAV injection surgical procedures and are performed under aseptic conditions.
	<ol>
		<li>Anesthetize the mouse with 3% isoflurane. 
		NOTE: This is the initial dose and reduce to 2% for the surgery later on. During imaging, an intraperitoneal (IP) injection of 130 mg ketamine/10 mg xylazine/kg body weight dissolved in ACSF is used for anesthesia.</li>
		<li>Position the mouse on the mouse stereotaxic device and fix the head with ear bars. Apply ophthalmic ointment to eyes. Use a heating pad to keep the body temperature of the mouse at 37 &#176;C throughout the surgery. 
		NOTE: All surgical tools need to be sterilized.</li>
		<li>Make an incision of 5-8 mm long in the midline of the scalp and remove a flap of skin using a pair of scissors.</li>
		<li>After the skull is exposed, perform 2.0-3.0 mm diameter craniotomy using a high-speed drill over the virus injected area (i.e., motor cortex or somatosensory cortex, Figure 1B). First, make four small holes, and then drill along in a circle connecting the holes. Then, lift and remove the bone with sharp scissors and remove. The exposed dura mater can be removed or kept intact for impantation of the cranial window.</li>
		<li>Place a glass coverslip of 3-5 mm in diameter carrying a transparent silicone over the craniotomy. Use a toothpick to push the cranial window gently onto the surface of the brain. Then, seal the edge with a small amount of silicone adhesive. 
		NOTES: Alternatively, instead of silicone disk, 1.2% low melting point agarose gel can be used between the cover glass and the brain tissue.</li>
		<li>Finally, seal the edges of the coverslip with dental cement. Take care to apply the cement slightly at the edge of the cranial window for strong bonding. Attach a custom-made metal head plate to the skull with dental cement. 
		NOTE: The metal plate is used to fix the head of the mouse to the stage of 2-P microscope during the imaging session (Figure 1C).</li>
		<li>Add 0.5 mL of ACSF solution over the coverslip on the cranial window for water immersion during imaging.</li>
		<li>Perform time-lapse in vivo 2-P imaging of mito-GCaMP5G in astrocytes and mito-GCaMP6s in neurons with 910 nm wavelength through the cranial window (Figure 1C)18,20,27. 
		NOTE: During imaging, mice are on the heating pad to maintain physiological temperature. The mice will be sacrificed immediately after the imaging session is completed.</li>
		<li>Label astrocytes in vivo with sulforhodamine 101 (SR101) when it is necessary to determine colocalization.
		<ol>
			<li>At the end of step 3.3.4, apply 100 &#181;L of 100 &#181;M SR101 in ACSF on the cortical surface for 1-5 min.</li>
			<li>Rinse the surface with ACSF to wash away the unbound SR101. Using 2-P imaging, co-labeling of mito-GCaMP5G and SR101 in astrocytes can be observed 45-60 min later (Figure 4A).</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

In this article, we provide a method and protocol for imaging mitochondrial Ca2+&#160;signals in astrocytes and neurons. We implemented mitochondria-targeting and cell type-specific strategies to express GECI GCaMP5G/6s. To target GCaMP5G/6s in mitochondria, we included a mitochondria-targeting sequence in the plasmids. To express GCaMP5G/6s in astrocytes and neurons in vivo, we inserted an astrocyte-specific promoter gfaABC1D and neuron-specific promoter CaMKII into the plasmids. Cell-type specific e...

Mitochondria are dynamic subcellular organelles and are considered as the cell powerhouses for energy production. On the other hand, mitochondria can take up Ca2+ to the matrix in response to local or cytosolic Ca2+ rises. Mitochondrial Ca2+ uptake affects mitochondrial function, including metabolic processes such as reactions in the tricarboxylic acid (TCA) cycle and oxidative phosphorylation, and regulates Ca2+-sensitive proteins under physiological conditions1,2,3,4. Mitochondrial Ca2+ overloading is also a determinant for cell death, including necrosis and apoptosis in various brain disorders5,6,7. It causes the opening of mitochondrial permeability transition pores (mPTPs) and the release of caspase cofactor, which initiate apoptotic cell death. Therefore, it is important to study mitochondrial Ca2+ dynamics and handling in living cells to understand cellular physiology and pathology better.
Mitochondria maintain matrix Ca2+ homeostasis through a balance between Ca2+ uptake and efflux. Mitochondrial Ca2+ uptake is mainly mediated by mitochondrial Ca2+ uniporters (MCUs), while mitochondrial Ca2+ efflux is mediated by the Na+-Ca2+-Li+ exchangers (NCLXs) and the H+/Ca2+ exchangers (mHCXs)8. The balance can be perturbed through the stimulation of G-protein coupled receptors (GPCRs)9. Mitochondrial Ca2+ homeostasis is also affected by mitochondrial buffering by the formation of insoluble xCa2+-xPO4x-xOH complexes8.
Intracellular and mitochondrial changes in Ca2+ concentration ([Ca2+]) can be evaluated by fluorescent or luminescent Ca2+ indicators. Ca2+ binding to indicators causes spectral modifications, allowing to recording of free cellular [Ca2+] in real-time in live cells. Two types of probes are currently available to monitor Ca2+ changes in cells: organic chemical indicators and genetically-encoded Ca2+ indicators (GECIs). Generally, different variants with different Ca2+ affinities (based on Kd), spectral properties (excitation and emission wavelengths), dynamic ranges, and sensitivities are available for the biological questions under investigation. Although many synthetic organic Ca2+ indicators have been used for cytosolic Ca2+ imaging, only a few can be selectively loaded in the mitochondrial matrix for mitochondrial Ca2+ imaging, with Rhod-2 being the most widely used (for reviews see10,11). However, Rhod-2 has a major drawback of leakage during long time-course experiments; in addition, it is partitioned between mitochondria, other organelles and the cytosol, making absolute measurements in different subcompartments difficult. In contrast, by using cell-type specific promoters and subcellular compartment targeting sequences, GECIs can be expressed in different cell types and subcellular compartments for cell- and compartment-specific Ca2+ imaging in vitro or in vivo. Single-wavelength fluorescence intensity-based GCaMP Ca2+ indicators have recently emerged as major GECIs12,13,14,15,16. In this article, we provide a protocol for mitochondria-targeting and cell-type specific expression of GCaMP5G and GCaMP6s (GCaMP5G/6s) in astrocytes and neurons, and imaging mitochondrial Ca2+ uptake in these cell types. Using this protocol, the expression of GCaMP6G/6s in individual mitochondria can be revealed, and Ca2+ uptake in single mitochondrial resolution can be achieved in astrocytes and neurons in vitro and in vivo.

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		</tr>
		<tr>
			<td>Cyanoacrylate glue</td>
			<td>World Precision Instruments</td>
			<td></td>
			<td>3M Vetbond Adhesive</td>
		</tr>
		<tr>
			<td>Dissecting stereomicroscope</td>
			<td>Nikon</td>
			<td>SMZ 2B</td>
			<td>Surgery</td>
		</tr>
		<tr>
			<td>Dumont forceps with fine tip</td>
			<td>Fine Science Tools</td>
			<td>11255-20</td>
			<td>for removal of dura</td>
		</tr>
		<tr>
			<td>Glass cover slips, 0.13-0.17 mm thick</td>
			<td>Fisher Scientific</td>
			<td>12-542A</td>
			<td>for cranial window cover</td>
		</tr>
		<tr>
			<td>High speed micro drill</td>
			<td>Fine Science Tools</td>
			<td>18000-17</td>
			<td>with bone polishing drill bit</td>
		</tr>
		<tr>
			<td>Injection syringe</td>
			<td>Hamilton</td>
			<td>2.5 ml</td>
			<td>for viral injection</td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>VEDCO</td>
			<td>NDC-50989-996-06</td>
			<td>100 mg/kg body weight</td>
		</tr>
		<tr>
			<td>Low melting point agarose</td>
			<td>Sigma-Aldrich</td>
			<td>A9793</td>
			<td>reducing movement artifacts</td>
		</tr>
		<tr>
			<td>Metal frame</td>
			<td>Custom-made</td>
			<td>see Fig 1</td>
			<td>for brain attachment to microscope stage</td>
		</tr>
		<tr>
			<td>MicroSyringe Pump Controller</td>
			<td>World Precision Instruments</td>
			<td>UMP3</td>
			<td>Injection speed controller</td>
		</tr>
		<tr>
			<td>Mouse stereotaxic device</td>
			<td>Stoelting</td>
			<td>51725</td>
			<td>for holding mice</td>
		</tr>
		<tr>
			<td>Perfusion chamber</td>
			<td>Warner Instruments</td>
			<td>64-0284</td>
			<td></td>
		</tr>
		<tr>
			<td>Persfusion system</td>
			<td>ALA Scientific Instruments</td>
			<td>ALA-VM8</td>
			<td></td>
		</tr>
		<tr>
			<td>Self-regulating heating pad</td>
			<td>Fine Science Tools</td>
			<td>21061</td>
			<td>to prevent hypothermia of mice</td>
		</tr>
		<tr>
			<td>Sulforhodamine 101</td>
			<td>Invitrogen</td>
			<td>S-359</td>
			<td>red fluorescent dye to label astrocytes</td>
		</tr>
		<tr>
			<td>Surgical scissors, 12 cm</td>
			<td>Fine Science Tools</td>
			<td>14002-12</td>
			<td>for dissection</td>
		</tr>
		<tr>
			<td>Trephine</td>
			<td>Fine Science Tools</td>
			<td>18004-23</td>
			<td>for clearing of material</td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>VEDCO</td>
			<td>NDC-50989-234-11</td>
			<td>10 mg/kg body weight</td>
		</tr>
	</tbody>
</table>

imaging mitochondrial ca2+ uptake in astrocytes and neurons using genetically encoded ca2+ indicators (gecis)

Mitochondrial Ca2+ plays a critical role in controlling cytosolic Ca2+ buffering, energy metabolism, and cellular signal transduction. Overloading of mitochondrial Ca2+ contributes to various pathological conditions, including neurodegeneration and apoptotic cell death in neurological diseases. Here we present a cell-type specific and mitochondria targeting molecular approach for mitochondrial Ca2+ imaging in astrocytes and neurons in vitro and in vivo. We constructed DNA plasmids encoding mitochondria-targeting genetically encoded Ca2+ indicators (GECIs) GCaMP5G or GCaMP6s (GCaMP5G/6s) with astrocyte- and neuron-specific promoters gfaABC1D and CaMKII and mitochondria-targeting sequence (mito-). For in vitro mitochondrial Ca2+ imaging, the plasmids were transfected in cultured astrocytes and neurons to express GCaMP5G/6s. For in vivo mitochondrial Ca2+ imaging, adeno-associated viral vectors (AAVs) were prepared and injected into the mouse brains to express GCaMP5G/6s in mitochondria in astrocytes and neurons. Our approach provides a useful means to image mitochondrial Ca2+ dynamics in astrocytes and neurons to study the relationship between cytosolic and mitochondrial Ca2+ signaling, as well as astrocyte-neuron interactions.

The aim of this study was to provide a methodology to image mitochondrial Ca2+ signals using GECIs in astrocytes and neurons in vitro and in vivo. Results of both in vitro and in vivo mitochondrial Ca2+ imaging are presented here.
In vitro mitochondrial Ca2+ signaling in cultured astrocytes and neurons 
Mitochondrial Ca2+ uptake in as...

Watch this Scientific Journal Video about Imaging Mitochondrial Ca2+ Uptake in Astrocytes and Neurons using Genetically Encoded Ca2+ Indicators GECIs at JoVE.com

Imaging Mitochondrial Ca2+ Uptake in Astrocytes and Neurons using Genetically Encoded Ca2+ Indicators GECIs

This proctocol aims to provide a method for in vitro and in vivo mitochondrial Ca2+ imaging in astrocytes and neurons.

Imaging Mitochondrial Ca2+ Uptake in Astrocytes and Neurons using Genetically Encoded Ca2+ Indicators (GECIs)

Mitochondrial Ca2+ plays a critical role in controlling cytosolic Ca2+ buffering, energy metabolism, and cellular signal transduction. Overloading of ...

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JoVE Journal

Neuroscience

4.0K Views.  University of Missouri-Columbia. This proctocol aims to provide a method for in vitro and in vivo mitochondrial Ca2+ imaging in astrocytes and neurons.

Video: Imaging Mitochondrial Ca2+ Uptake in Astrocytes and Neurons using Genetically Encoded Ca2+ Indicators GECIs

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

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

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

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

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate retina. Two classes of retinal neurons, photoreceptors and bipolar cells, accomplish this by using ribbon-type active zones, which enable sustained and high-throughput neurotransmitter release over long time periods. ON-type mixed bipolar cell (Mb) terminals in the goldfish retina, which depolarize to light stimuli and receive mixed rod and cone photoreceptor input, are suitable for the study of ribbon-type synapses both due to their large size (~10-12 &mu;m diameter) and to their numerous lateral and reciprocal synaptic connections with amacrine cell dendrites. Direct access to Mb bipolar cell terminals in goldfish retinal slices with the patch-clamp technique allows the measurement of presynaptic Ca2+ currents, membrane capacitance changes, and reciprocal synaptic feedback inhibition mediated by GABAA and GABAC receptors expressed on the terminals. Presynaptic membrane capacitance measurements of exocytosis allow one to study the short-term plasticity of excitatory neurotransmitter release 14,15. In addition, short-term and long-term plasticity of inhibitory neurotransmitter release from amacrine cells can also be investigated by recordings of reciprocal feedback inhibition arriving at the Mb terminal 21. Over short periods of time (e.g. ~10 s), GABAergic reciprocal feedback inhibition from amacrine cells undergoes paired-pulse depression via GABA vesicle pool depletion 11. The synaptic dynamics of retinal microcircuits in the inner plexiform layer of the retina can thus be directly studied.
The brain-slice technique was introduced more than 40 years ago but is still very useful for the investigation of the electrical properties of neurons, both at the single cell soma, single dendrite or axon, and microcircuit synaptic level 19. Tissues that are too small to be glued directly onto the slicing chamber are often first embedded in agar (or placed onto a filter paper) and then sliced 20, 23, 18, 9. In this video, we employ the pre-embedding agar technique using goldfish retina. Some of the giant bipolar cell terminals in our slices of goldfish retina are axotomized (axon-cut) during the slicing procedure. This allows us to isolate single presynaptic nerve terminal inputs, because recording from axotomized terminals excludes the signals from the soma-dendritic compartment. Alternatively, one can also record from intact Mb bipolar cells, by recording from terminals attached to axons that have not been cut during the slicing procedure. Overall, use of this experimental protocol will aid in studies of retinal synaptic physiology, microcircuit functional analysis, and synaptic transmission at ribbon synapses.

Patch-clamp Capacitance Measurements and Ca2+ Imaging at Single Nerve Terminals in Retinal Slices

Here we describe a protocol for the preparation of agar-embedded retinal slices that are suitable for electrophysiology and Ca2+ imaging. This method allows one to study ribbon-type synapses in retinal microcircuits using direct patch-clamp recordings of single presynaptic nerve terminals.

Visual stimuli are detected and conveyed over a wide dynamic range of light intensities and frequency changes by specialized neurons in the vertebrate ...

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within the same species 1,2. These intraspecies signals, the pheromones, as well as signals from some predators 3, activate the vomeronasal sensory neurons (VSNs) with high levels of specificity and sensitivity 4. At least three distinct families of G-protein coupled receptors, V1R, V2R and FPR 5-14, are expressed in VNO neurons to mediate the detection of the chemosensory cues. To understand how pheromone information is encoded by the VNO, it is critical to analyze the response profiles of individual VSNs to various stimuli and identify the specific receptors that mediate these responses.
The neuroepithelia of VNO are enclosed in a pair of vomer bones. The semi-blind tubular structure of VNO has one open end (the vomeronasal duct) connecting to the nasal cavity. VSNs extend their dendrites to the lumen part of the VNO, where the pheromone cues are in contact with the receptors expressed at the dendritic knobs. The cell bodies of the VSNs form pseudo-stratified layers with V1R and V2R expressed in the apical and basal layers respectively 6-8. Several techniques have been utilized to monitor responses of VSNs to sensory stimuli 4,12,15-19. Among these techniques, acute slice preparation offers several advantages. First, compared to dissociated VSNs 3,17, slice preparations maintain the neurons in their native morphology and the dendrites of the cells stay relatively intact. Second, the cell bodies of the VSNs are easily accessible in coronal slice of the VNO to allow electrophysiology studies and imaging experiments as compared to whole epithelium and whole-mount preparations 12,20. Third, this method can be combined with molecular cloning techniques to allow receptor identification.
Sensory stimulation elicits strong Ca2+ influx in VSNs that is indicative of receptor activation 4,21. We thus develop transgenic mice that express G-CaMP2 in the olfactory sensory neurons, including the VSNs 15,22. The sensitivity and the genetic nature of the probe greatly facilitate Ca2+ imaging experiments. This method has eliminated the dye loading process used in previous studies 4,21. We also employ a ligand delivery system that enables application of various stimuli to the VNO slices. The combination of the two techniques allows us to monitor multiple neurons simultaneously in response to large numbers of stimuli. Finally, we have established a semi-automated analysis pipeline to assist image processing.

The vomeronasal organ (VNO) detects intraspecies chemical signals that convey social and reproductive information. We have performed Ca2+ imaging experiments using transgenic mice expressing G-CaMP2 in VNO tissue. This approach allows us to analyze the complicated response patterns of the vomeronasal neurons to large numbers of pheromone stimuli.

The vomeronasal organ (VNO) detects chemosensory signals that carry information about the social, sexual and reproductive status of the individuals within ...

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with enhanced [Ca2+]i signals. It has been proposed that astrocytes in turn regulate neurons and blood vessels through calcium-dependent mechanisms, such as the release of signaling molecules. However, [Ca2+]i imaging in entire astrocytes has only recently become feasible with genetically encoded calcium indicators (GECIs) such as the GCaMP series. The use of GECIs in astrocytes now provides opportunities to study astrocyte [Ca2+]i signals in detail within model microcircuits such as the striatum, which is the largest nucleus of the basal ganglia. In the present report, detailed surgical methods to express GECIs in astrocytes in vivo, and confocal imaging approaches to record [Ca2+]i signals in striatal astrocytes in situ, are described. We highlight precautions, necessary controls and tests to determine if GECI expression is selective for astrocytes and to evaluate signs of overt astrocyte reactivity. We also describe brain slice and imaging conditions in detail that permit reliable [Ca2+]i imaging in striatal astrocytes in situ. The use of these approaches revealed the entire territories of single striatal astrocytes and spontaneous [Ca2+]i signals within their somata, branches and branchlets. The further use and expansion of these approaches in the striatum will allow for the detailed study of astrocyte [Ca2+]i signals in the striatal microcircuitry.

Imaging Intracellular Ca2+ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

The properties and functions of astrocyte intracellular Ca2+ signals in the striatum remain incompletely explored. We describe methods to express genetically encoded calcium indicators in striatal astrocytes using adeno-associated viruses of serotype 2/5 (AAV2/5), as well as procedures to reliably image Ca2+ signals within striatal astrocytes in situ.

Astrocytes display spontaneous intracellular Ca2+ concentration fluctuations ([Ca2+]i) and in several settings respond to neuronal excitation with ...

In Situ Ca2+ Imaging of the Enteric Nervous System

Reflex behaviors of the intestine are controlled by the enteric nervous system (ENS). The ENS is an integrative network of neurons and glia in two ganglionated plexuses housed in the gut wall. Enteric neurons and enteric glia are the only cell types within the enteric ganglia. The activity of enteric neurons and glia is responsible for coordinating intestinal functions. This protocol describes methods for observing the activity of neurons and glia within the intact ENS by imaging intracellular calcium (Ca2+) transients with fluorescent indicator dyes. Our technical discussion focuses on methods for Ca2+ imaging in whole-mount preparations of the myenteric plexus from the rodent bowel. Bulk loading of ENS whole-mounts with a high-affinity Ca2+ indicator such as Fluo-4 permits measurements of Ca2+ responses in individual neurons or glial cells. These responses can be evoked repeatedly and reliably, which permits quantitative studies using pharmacological tools. Ca2+ responses in cells of the ENS are recorded using a fluorescence microscope equipped with a cooled charge-coupled device (CCD) camera. Fluorescence measurements obtained using Ca2+ imaging in whole-mount preparations offer a straightforward means of characterizing the mechanisms and potential functional consequences of Ca2+ responses in enteric neurons and glial cells.

In Situ Ca2+ Imaging of the Enteric Nervous System

The enteric nervous system (ENS) is a network of neurons and glia located in the gut wall that controls intestinal reflexes. This protocol describes methods for recording the activity of enteric neurons and glia in live preparations of ENS using Ca2+ imaging.

Reflex behaviors of the intestine are controlled by the enteric nervous system (ENS). The ENS is an integrative network of neurons and glia in two ...

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to blindness in many retinal diseases. Calcium (Ca2+), a key second messenger in photoreceptor signaling and metabolism, has been proposed to be indirectly linked with photoreceptor degeneration in various animal models. Systematically studying these aspects of cone physiology and pathophysiology has been hampered by the difficulties of electrically recording from these small cells, in particular in the mouse where the retina is dominated by rod photoreceptors. To circumvent this issue, we established a two-photon Ca2+ imaging protocol using a transgenic mouse line that expresses the genetically encoded Ca2+ biosensor TN-XL exclusively in cones and can be crossbred with mouse models for photoreceptor degeneration. The protocol described here involves preparing vertical sections (&#8220;slices&#8221;) of retinas from mice and optical imaging of light stimulus-evoked changes in cone Ca2+ level. The protocol also allows &#8220;in-slice measurement&#8221; of absolute Ca2+ concentrations; as the recordings can be followed by calibration. This protocol enables studies into functional cone properties and is expected to contribute to the understanding of cone Ca2+ signaling as well as the potential involvement of Ca2+ in photoreceptor death and retinal degeneration.

Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

We describe a protocol to monitor Ca2+ dynamics in the axon terminals of cone photoreceptors using an ex-vivo&#160;slice preparation of the mouse retina. This protocol allows comprehensive studies of cone Ca2+ signaling in an important mammalian model system, the mouse.

Retinal cone photoreceptors (cones) serve daylight vision and are the basis of color discrimination. They are subject to degeneration, often leading to ...

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate goal is to image neuronal activity in vivo, one must be able to image activity of a single cell to ensure successful in vivo preparations. This procedure will describe how to image membrane potential in a single cell to provide a foundation to eventually image in vivo. Here we describe methods for imaging GEVIs consisting of a voltage-sensing domain fused to either a single fluorescent protein (FP) or two fluorescent proteins capable of F&#246;rster resonance energy transfer (FRET) in vitro. Using an image splitter enables the projection of images created by two different wavelengths onto the same charge-coupled device (CCD) camera simultaneously. The image splitter positions a second filter cube in the light path. This second filter cube consists of a dichroic and two emission filters to separate the donor and acceptor fluorescent wavelengths depending on the FPs of the GEVI. This setup enables the simultaneous recording of both the acceptor and donor fluorescent partners while the membrane potential is manipulated via whole cell patch clamp configuration. When using a GEVI consisting of a single FP, the second filter cube can be removed allowing the mirrors in the image splitter to project a single image onto the CCD camera.

A method for imaging changes in membrane potential using genetically encoded voltage indicators is described.

Genetically encoded voltage indicators (GEVIs) have improved to the point where they are beginning to be useful for in vivo recordings. While the ultimate ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Mitochondrial Ca2+ Retention Capacity Assay and Ca2+-triggered Mitochondrial Swelling Assay

The production of ATP by oxidative phosphorylation is the primary function of mitochondria. Mitochondria in higher eukaryotes also participate in cytosolic Ca2+ buffering, and the ATP production in mitochondrial can be mediated by intramitochondrial free Ca2+ concentration. Ca2+ retention capacity can be regarded as the capability of mitochondria to retain calcium in the mitochondrial matrix. Accumulated intracellular Ca2+ leads to the permeability of the inner mitochondrial membrane, termed the opening of mitochondrial permeability transition pore (mPTP), which leads to the leakage of molecules with a molecular weight less than 1.5 kDa. Ca2+-triggered mitochondria swelling is used to indicate the mPTP opening. Here, we describe two assays to examine the Ca2+ retention capacity and Ca2+-triggered mitochondrial swelling in isolated mitochondria. After certain amounts of Ca2+ are added, all steps can be completed in one day and recorded by a microplate reader. Thus, these two simple and effective assays can be adopted to assess the Ca2+-related mitochondrial functions.

Mitochondrial Ca2+ Retention Capacity Assay and Ca2+-triggered Mitochondrial Swelling Assay

This protocol aims to describe a method to examine the Ca2+ retention capacity and Ca2+- triggered mitochondrial swelling of isolated mitochondria of SH-SY5Y cells step-by-step.

Imaging Mitochondrial Ca²⁺ Uptake in Astrocytes and Neurons using Genetically Encoded Ca²⁺ Indicators (GECIs)

Summary

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Chapters in this video

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

Patch-clamp Capacitance Measurements and Ca²⁺ Imaging at Single Nerve Terminals in Retinal Slices

Imaging Neuronal Responses in Slice Preparations of Vomeronasal Organ Expressing a Genetically Encoded Calcium Sensor

Imaging Intracellular Ca²⁺ Signals in Striatal Astrocytes from Adult Mice Using Genetically-encoded Calcium Indicators

In Situ Ca²⁺ Imaging of the Enteric Nervous System

Imaging Ca²⁺ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina

Fluorescence and Bioluminescence Imaging of Subcellular Ca²⁺ in Aged Hippocampal Neurons

Imaging Membrane Potential with Two Types of Genetically Encoded Fluorescent Voltage Sensors

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Mitochondrial Ca²⁺ Retention Capacity Assay and Ca²⁺-triggered Mitochondrial Swelling Assay