This work was supported by the Deutsche Forschungsgemeinschaft (DFG) (Werner Reichardt Centre for Integrative Neuroscience T&#252;bingen, EXC 307 to T.E. and T.S.; KFO 134 to B.W. and F.P.D.), the German Federal Ministry of Education and Research (BMBF) (BCCN T&#252;bingen, FKZ 01GQ1002 to T.E and T.B.), and the European Union (DRUGSFORD; HEALTH-F2-2012-304963 to M.K. and F.P.D).

All animal procedures were carried out adhering to the guidelines and laws for animal protection determined by German Federal Government and approved by the institutional animal welfare committee of the University of T&#252;bingen.
1. Animal Models
<ol>
	<li>HR2.1:TN-XL Ca2+ biosensor mouse
	<ol>
		<li>Use the transgenic HR2.1:TN-XL mouse line expressing the Ca2+ biosensor TN-XL selectively in cone photoreceptors10. Use 3 - 6 weeks old mice of either sex raised with a standard 12 hrs day/night rhythm.</li>
	</ol>
	</li>
</ol>
2. Retinal Dissection
<ol>
	<li>Physiological solution
	<ol>
		<li>Prepare freshly 2 L&#160;of extracellular solution, containing 125 mM NaCl, 2.5 mM KCl, 1 mM MgCl2, 1.25 mM NaH2PO4, 26 mM NaHCO3, 0.5 mM L-glutamine, and 20 mM (+) glucose. Mix until all ingredients dissolve.</li>
		<li>&#34;Bubble&#34;&#160;extracellular solution with carboxygen (95% O2, 5% CO2) for 5 - 10 min. Add CaCl2 to reach a concentration of 2 mM.</li>
		<li>After bubbling the extracellular solution, measure its pH. The pH should be 7.4, otherwise it is likely that mistakes have been made during the preparation of the solution. Keep bubbling rate and solution temperature constant throughout experiment to ensure constant pH.</li>
		<li>Separate the stock into 2 flasks, one for retina dissection and one for the recording setup (perfusion).</li>
	</ol>
	</li>
	<li>Eye enucleation and isolation of retina (5 - 10 min)
	<ol>
		<li>Maintain a dim red illumination in the working area for enucleation. Use LEDs with a peak wavelength of 650 nm (or longer) to avoid bleaching of cones during dissection.</li>
		<li>Dark-adapt the mouse for 2 hrs by putting its cage into a well-ventilated, lightproof box to ensure that cones are fully sensitized at time of recordings.</li>
		<li>Anaesthetize the mouse under a laboratory hood with isoflurane (5%) using a vaporizer and a gas-tight container that holds a standard mouse cage. Carefully follow the manufacturer&#180;s instructions when handling the vaporizer. Use gloves and face mask to reduce exposure to allergens.</li>
		<li>Use a binocular microscope with a 10 - 40X&#160;magnification for dissection.</li>
		<li>Sacrifice the anaesthetized mouse by decapitation.</li>
		<li>For orientation, mark the top of each eye (= dorsal) with a waterproof pen. Keep track on whether using left or right eye.</li>
		<li>Remove the eye carefully using curved scissors by cutting the optic nerve behind the eyeball. For dissection, transfer the eyeball to a Petri dish containing freshly carboxygenated extracellular solution. For transferring the eyeball, hold it by the optic nerve stump.</li>
		<li>Pierce the eye at any point along the border between the cornea and the sclera (i.e., at the ora serrata) with a sharp injection needle.</li>
		<li>Hold the eye at the sclera with a pair of forceps and gently insert one scissors blade into the hole made in the previous step. Cut along the ora serrata to separate the anterior part of the eye (cornea, lens, vitreous body) from the posterior part (eyecup). Cut the eyecup radially (towards the optic disc) at the position of the pen mark to indicate the dorsal position on the retina. 
		Note: Prepared this way, the eyecups can be kept in constantly carboxygenated solution for later use.</li>
		<li>Turn the eyecup in the Petri dish such that the dorsal cut points away from oneself. Grab the sclera on the left and the right side of the eyecup with a pair of forceps each by inserting the forceps tips between retina and sclera.</li>
		<li>Detach the retina by gently inverting the scleral part of the eyecup. Finally, cut the optic nerve using micro dissecting scissors to free the retina from the sclera. 
		Note: This is a critical step. Avoid damage to the retina (e.g., by touching and squeezing it with forceps).</li>
		<li>Hold one of the edges of the retina with a pair of forceps and gently remove debris and vitreous body from the retinal surface using a second pair of forceps. When the retinal surface is clean, use micro dissecting scissors to make three additional shorter, radial cuts (approximately every 90&#176;). These cuts allow one to carefully flatten the retina with the photoreceptor side facing down in the Petri dish (Figure 1A).</li>
	</ol>
	</li>
	<li>Slice preparation (10 - 15 min)
	<ol>
		<li>Prepare beforehand nitrocellulose filter membranes for retinal slicing and glass cover-slips for mounting slices and transferring them to the recording chamber. Cut nitrocellulose filter membrane into approximately 10 x 5 mm sized rectangular pieces using scissors. Cut coverslips into approximately 10 x 5 mm sized rectangular pieces using a glasscutter.</li>
		<li>Slowly immerse a glass slide into the extracellular solution close to tissue. Gently, pull the retina onto the glass slide with the ganglion cell side up by grabbing it at the very edge using a pair of forceps. This process reduces mechanical damage and folding of tissue.</li>
		<li>Choose the region of the retina that is related to the respective research question (see also Discussion). Remember, the long cut made in step 2.2.9 marks the dorsal retinal half (Figure 1A). Cut a rectangular, approximately 1 x 2 mm sized piece out of the selected retinal region using a curved scalpel blade. Wipe off excess solution around the tissue.</li>
		<li>Place the filter membrane on top of the piece of retina such that the ganglion cell side adheres to the membrane. Immediately, add a drop of extracellular medium onto the membrane to firmly attach the tissue to the membrane.</li>
		<li>Transfer the membrane-mounted retinal tissue to the slicing chamber containing fresh extracellular solution.</li>
		<li>Cut retina into vertical slices of 200 &#181;m thickness (Figure 1B) using a fresh razor blade attached to a tissue chopper23. Change blade for every retinal piece. 
		Note: The razor blade needs to be perfectly aligned with the surface of the slicing chamber bottom such that the whole membrane splits simultaneously, as indicated by a &#8220;clicking&#8221; sound when cutting &#8211; otherwise the blade may bend and damage the slice.</li>
		<li>Glue a single membrane-mounted slice to a glass cover-slip by applying high vacuum grease to the membrane ends only (Figure 1C). Keep the glass surface below the retina free from grease.</li>
		<li>Keep coverslip-mounted slices covered by a drop of extracellular solution in the holding chamber &#8211; a closed and lightproof container, e.g., a Petri dish with the lid covered with aluminium foil &#8211; under a carboxygen atmosphere at RT. Introduce carboxygen by bubbling a small water reservoir to keep the atmosphere in the holding chamber humidified; this prevents the slices from drying out.</li>
		<li>Allow slices to rest in the holding chamber for 10 - 15 min before moving them (one by one) to the recording chamber. Slices can be maintained up to 4-5 hrs in holding chamber at RT (~21 &#176;C).</li>
	</ol>
	</li>
</ol>
3. Two-photon Ca2+&#160;Imaging
<ol>
	<li>Two-photon microscopy
	<ol>
		<li>Use a Movable Objective Microscope (MOM)-type two-photon microscope. Both MOM design and imaging procedures were described earlier24, for details see also10,21,25&#160;(for sources and companies, see Table 1). 
		Note: Any upright two-photon microscope that fulfils the following minimal requirements can be used: It has to be equipped with (a) a pulsed laser tuneable to ~860 nm, (b) a minimum of two simultaneously acquired fluorescence channels, (c) filters for eCFP and citrine fluorescence, (d) a light stimulator (for possible designs, see24,26), and (e) software that allows recording time-lapsed image sequences at a frame rate sufficient to resolve the Ca2+ signals of interest.</li>
		<li>Start the two-photon imaging system as indicated by the manufacturer. Strictly follow the facility&#8217;s laser safety guidelines. Start the laser and tune it to ~860 nm.</li>
		<li>Transfer a slice from the holding chamber to the recording chamber and immediately start perfusing with carboxygenated extracellular solution. Maintain a perfusion flow rate of 2 ml/min and a temperature of 37 &#176;C in the recording chamber.</li>
		<li>Use a 20X 0.95 NA water immersion objective. If available, use a CCD camera in combination with an infra-red LED below recording chamber to locate the retinal slice (Figure 2). Otherwise locate slice using two-photon imaging (see 3.1.5).</li>
		<li>Switch to two-photon imaging to view biosensor expression. Turn on the two detection channels for fluorescence imaging of eCFP and citrine.</li>
		<li>Use the image acquisition software that controls the two-photon microscope to scan and select a row of cone terminals for recording (Figure 3A). Set image acquisition to 128 x 16 pixel images (31.25 Hz) or a similar configuration. Restrict scanned area to cone terminals to avoid bleaching of photopigments in outer segments. 
		Note: For the TN-XL calcium sensor (&#964; = ~0.6 sec for Ca2+ binding, &#964; = ~0.2 sec for Ca2+ unbinding; see16) a minimum frame rate of ~8 Hz is recommended.</li>
	</ol>
	</li>
	<li>Light stimulation and recording
	<ol>
		<li>Use a sub-stage full-field light stimulator as described elsewhere10,21,26 to perform the following steps. 
		Note: A simple solution for a full-field light stimulator is to use two band-pass-filtered LEDs (e.g., &#8220;blue&#8221;: 360 BP 12, &#8220;green&#8221;: 578 BP 10) that match the wavelength sensitivities of mouse cones but at the same time do not overlap with the filters used for fluorescence detection (cf. 3.2.4). The light from the LEDs is focused by the condenser through the bottom of the recording chamber (Figure 2). For details on this stimulator design, its calibration, the light intensities used and the evoked cone photo-isomerisation rates, see10,21,26.</li>
		<li>Turn on the laser and allow cones to adapt to the scanning laser and the stimulus background light (20 - 30 sec, see also Potential Pitfalls) before presenting light stimuli (see 3.2.3) or applying pharmacological agents.</li>
		<li>Start presentation of arbitrary stimuli (e.g., flashes of light, as shown in Figure 3B,C). In the case of the stimulator described in step 3.2.1, generate the stimuli by modulating the intensity of the two LEDs over time using a microprocessor board controlled by customized software (for details, see10,21,26).</li>
		<li>Start recording the two fluorescence channels (e.g., at 483 nm for &#8220;blue&#8221; FRET-donor eCFP and at 535 nm for &#8220;yellow&#8221; FRET-acceptor; for specs see Table 1) simultaneously using the respective image acquisition software (see also 3.1.6). For example, in the case of light flashes (Figure 3B,C), record at least 8 - 10 stimulus presentations (trials).</li>
	</ol>
	</li>
	<li>&#8220;In-slice&#8221; Ca2+ calibration
	<ol>
		<li>Record Ca2+ signals in cone terminals (as described in 3.1.6 and 3.2.4) and extract the baseline Ca2+ level in a set of cones (as described in 3.4).</li>
		<li>Switch to &#8220;Ca2+ free&#8221; extracellular medium with 5&#160;&#181;M ionomycin (dissolved in DMSO) and 10 mM EGTA (or, alternatively, BAPTA) added. Record cone terminals again every 5 min to determine the minimal fluorescence ratio (Rmin), i.e., the ratio in the (near) absence of intracellular Ca2+. This may take anywhere between 15 - 30 min. 
		Note: A perfusion system is required that allows switching between different reservoirs.</li>
		<li>Switch to extracellular medium with 5 &#181;M ionomycin (dissolved in DMSO) and 2.5 mM Ca2+. After an incubation time of 5 min, record cones again every 5 min to measure the maximal fluorescence ratio (Rmax), i.e., the ratio in the presence of saturating concentrations of intracellular Ca2+. Following ionomycin application, it may take between 3 - 15 min for the Ca2+ ratio to become stable. 
		Note: Exposing the slices to high Ca2+ and ionomycin for a longer period will eventually damage the cells. Include only cells in the analysis that retain their normal morphology. Concentrations (i.e., EGTA, ionomycin) may need to be adapted. For more details on the Ca2+ calibration protocol, see13,17.</li>
	</ol>
	</li>
	<li>Data analysis 
	&#160;Note: Use an image processing and data analysis software package compatible with the respective microscope software and capable of running custom analysis scripts.
	<ol>
		<li>Load an imaging data file and draw regions of interest (ROIs) around each cone terminal (Figure 3A). For each frame, average fluorescence intensity within the ROI and subtract background fluorescence, before calculating the ratio (R) between FRET-acceptor (citrine) and donor (eCFP) fluorescence (R=FA/FD; Figure 3B,C). This ratio is typically used as a proxy for relative intracellular Ca2+ concentration, but after calibration (see 3.3), also absolute concentrations can be estimated (see 3.4.3). 
		Note: The pedicles can be recognized by their location in the outer plexiform layer and their morphology (somewhat flattened &#8220;blobs&#8221; smaller than the cone somata, located at the end of the cone axon).</li>
		<li>Average stimulus trials (Figure 4A). Extract response parameters from averaged traces (Figure 4B), such as Ca2+ baseline level (Rbase) prior to light stimulation, peak amplitude (Ramp), and area under the curve (RA) as measures for response size. Also, extract response kinetics from averaged traces, such as response rise (trise) and decay time (tdecay) (= respective time intervals between 20% and 80% of Ramp; see illustration in Figure 4B). For more details on how to determine these parameters, see10.</li>
		<li>Calculate an estimate of the absolute cone Ca2+ concentration based on the measurements from steps 3.3.1-3.3.3. Use the ratios for minimal and maximal Ca2+ concentration, Rmin and Rmax, respectively, the donor fluorescence in the Ca2+-bound (FD,Ca-bound) and -unbound (FD,Ca-free) state, as well as the in-vivo dissociation constant (Kd&#160;= 0.77, see16) for TN-XL with the following equation (analogue to27):</li>
	</ol>
	</li>
</ol>
<img alt="Equation 1" src="/files/ftp_upload/52588/52588eq1.jpg" />

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The authors have nothing to disclose.

Pre-existing protocols using electrophysiological single-cell recordings or Ca2+ imaging with synthetic fluorescent indicators struggle to record the Ca2+ dynamics in mouse cone photoreceptors for a number of technical reasons (see Introduction). The protocol described here allows the measurement of Ca2+ signals and even absolute Ca2+ levels in individual, identified mouse cone terminals in an efficient and relatively simple way.
This protocol has already been successfully used in three studies addressing different aspects of cone function in healthy mouse retina. In the first study10, the HR2.1:TN-XL mouse was characterized using immunohistochemistry, ERG recordings, two-photon Ca2+ imaging, and pharmacology, showing that the cone-specific expression of the Ca2+ biosensor does not hamper cone anatomy and function. In the second study21, chromatic and achromatic response properties of mouse cones were mapped across the retina, demonstrating striking differences in cone function between the &#8220;green&#8221; opsin-dominated dorsal and the &#8220;blue&#8221; opsin-dominated ventral mouse retina. These regional differences in cone properties matched the differential contrast distribution in the natural environment (i.e., sky vs. ground), suggesting that the different spectral types of mouse cones provide for (near) optimal sampling of achromatic contrasts and, thus, may offer an evolutionary advantage. In the third study22 the reciprocal feedback that cone axon terminals receive from horizontal cells was investigated. As all proposed horizontal cell feedback mechanisms act on voltage-gated Ca2+ channels in the cone axon terminals28, cone terminal Ca2+ can serve as a proxy for horizontal cell-to-cone interactions. The study by Kemmler and coworkers22 supports the view that horizontal cells use a complex feedback system comprising several mechanisms to control photoreceptor glutamate release.
These studies illustrate the versatility of the described protocol and show that it can be adapted to a wide range of questions concerning cone function and its synaptic circuits. In addition, the protocol enables studying local Ca2+ signaling in the different cone compartments, towards a better comprehension of cone physiology. Such knowledge is important to understand pathophysiological processes in degenerating cones, to eventually allow for the rational development of potential therapeutic approaches, in particular for degenerative diseases affecting cones.
In the HR2.1:TN-XL mouse line, the Ca2+ biosensor is expressed throughout the cone, with the exception of the outer segment. This provides an opportunity for direct and ratiometric assessment of Ca2+ dynamics in different cone compartments. Since changes in Ca2+ currents in the outer segment are reflected in terminals via the membrane potential and the resulting activation of voltage-gated Ca2+ channels, processes in the outer segment can be observed indirectly.
Potential pitfalls:
The dissection of the retina is a critical step: In the mouse, the retina separates from the eyecup usually between photoreceptor outer segments and pigment epithelium. Therefore, the light-sensitive photoreceptor outer segments of the isolated retina are exposed and extremely sensitive to mechanical damage. Great care must be taken not to damage by touching the photoreceptor side with tools or by moving the tissue sideways on an adhesive surface (e.g., a filter membrane).
High-quality retinal slices can be recognized under the microscope by their clean cutting surface and by a well-organized photoreceptor layer with clearly defined outer segments. Functional assessment of slice quality can be quickly done by flashing bright light stimuli and determining the percentage of responsive cones (e.g., in a field of view with 10 - 20 cones). Here, response quality should be evaluated by calculating the signal-to-noise ratio (S/N) (amplitude of baseline noise before the light stimulus vs. amplitude of the light response); a S/N of 2 - 3 should be considered as the minimum threshold. Typically, we discard slices with less than 50% responsive cones. Also slices with cones that display excessive spontaneous spiking behaviour (see Figure&#160;4&#160;in10) should be discarded.
Slices in the recording chamber that meet the aforementioned anatomical and functional criteria show consistent responses for 1 - 2 hr (for details on response consistency, see10). Because slices survive for hours in the holding chamber, a successful experiment can last up to 6 hr. It is noteworthy that there are some restrictions with respect to studying long-ranging spatial interactions between cones and horizontal cells, as slicing inevitably severs lateral connections in retinal networks. However, increasing the thickness of retinal slices to 300&#160;&#181;m ameliorates this issue22.
The use of vertical retinal slices avoids scanning of light-sensitive cone outer segments by the excitation laser and, thus, largely prevents opsin bleaching (for extensive discussion, see10,21). Nevertheless, the recorded Ca2+ signals not only depend on the light stimuli, but also get affected by a background illumination component generated by the scanning excitation laser. In fact, the effective background illumination in such two-photon imaging experiments is a combination of scattered laser light, fluorescent light emitted by the recorded cells, and any LED stimulus background component. Therefore, cones should be allowed to adapt for at least 20 - 30 s to laser scanning (with the background component of the light stimulus turned on) prior to recording.
Advantages and applications:
While some applications for this cone Ca2+-imaging protocol have been described above10,21,22, other applications can be envisioned: Pharmacological studies in combination with cone Ca2+ imaging may validate Ca2+ signaling pathways in cones and could be used to test efficacy and potency of drugs targeting different players in Ca2+-signaling10. However, a key application may be to study diseases affecting cone function. Many retinal degeneration mouse models mimicking human diseases are available. For instance, the cone photoreceptor loss 1 (cpfl1) mouse is a primary cone degeneration model suffering from a Pde6c mutation29. Conversely, the rod degeneration 1 (rd1) mouse suffers from a Pde6b mutation. While this causes primary rod photoreceptor degeneration30, once rd1 rod loss is completed, secondary cone degeneration sets in1. Crossbreeding these animals with the HR2.1:TN-XL line will allow studying and comparing Ca2+ dynamics in both primary and secondary cone degeneration and is likely to provide valuable insights into the role of Ca2+ during cone cell death. Also, pharmacologically induced cone degeneration &#8211; for instance using selective PDE6 inhibitors &#8211; may serve to identify the downstream mechanisms of cone degeneration10,29,31.
In summary, the protocol described here allows measuring Ca2+ in subcellular compartments of mouse cone photoreceptors and presents great opportunities to uncover cone physiology under a wide range of physiological and pathophysiological conditions. Additionally, this protocol may be used for the screening of pharmacological agents designed to interfere with cone Ca2+-signaling and thus help to establish new therapies for cone diseases.

Vision begins with the light-induced activation of the phototransduction cascade in retinal photoreceptors. Rod photoreceptors allow vision at low light levels, whereas cone photoreceptors mediate colour and high-resolution daylight vision. Many photoreceptor-specific genes are susceptible to mutations that lead to degeneration of these cells. A number of molecular markers associated with photoreceptor loss have been identified1, but so far the detailed molecular mechanisms and the sequence of events remain unclear. Altered Ca2+ homeostasis was speculated to be a trigger of photoreceptor cell death, a hypothesis supported by the up-regulation of the activity of Ca2+-dependent calpain-type proteases during the degeneration process2,3. However, to date, this hypothesis is not backed by physiological measurements of Ca2+. Inconsistencies in several studies on the effect of Ca2+ channel blockers in retinal diseases further challenged the involvement of Ca2+ in cell death4-6, calling for methods to directly assess Ca2+ in mammalian cone photoreceptors.
Previously, most electrical recording and Ca2+ imaging studies have been performed in amphibian and reptilian models because of the easier access to cones7-9. However, mammalian photoreceptor physiology may be different from that of non-mammals10 and especially, in the context of human hereditary retinal degeneration, a better understanding of mammalian photoreceptor physiology is a key to the development of innovative treatments. Many mouse models mimicking human retinal diseases are available but little is known about Ca2+ dynamics in mouse cones11. Electrical techniques are not suitable for high-throughput recordings from cones, in particular not in mice, where rods (~97%) significantly outnumber cones (~3%)12. Moreover, sensitive electrophysiological techniques such as whole-cell patch-clamp recordings disturb the intracellular milieu and provide data on Ca2+ currents but not on absolute intracellular Ca2+ concentrations. Hence, despite the lower temporal resolution, imaging is the method of choice when addressing questions about cone Ca2+ dynamics. The key issue with imaging is how to selectively label the cones with a fluorescent Ca2+ indicator dye. Proper compartmentalization and cell specificity is hard to achieve by &#8220;bulk-loading&#8221; synthetic Ca2+ indicator dyes into the tissue. As a consequence, labelled cones, rods13,14, and M&#252;ller glial cells cannot be reliably distinguished. Also, synthetic dyes tend to leak out of the cells, preventing prolonged recordings under consistent conditions. Furthermore, loading synthetic Ca2+ indicators in their AM-ester form is problematic, as it requires detergents (e.g., DMSO) and generates formaldehyde15. For absolute Ca2+ measurements, ratiometric indicators are mandatory. However, the best currently available synthetic ratiometric indicator Fura-2 requires excitation light in the range of 700 to 760 nm (for two-photon excitation), which, depending on its intensity, can in itself stimulate the cones, and thus impede studies of cone Ca2+ dynamics under physiological illumination conditions.
Unlike synthetic dyes, genetically-encoded Ca2+ indicators can be expressed in a cell type-selective manner. They do not leak out of cells, and therefore, if bleaching is avoided, prolonged and reliable ratiometric measurements are possible. Cell type-selective expression of Ca2+ biosensors, when combined with two-photon microscopy, represents a powerful tool to assess and study subcellular Ca2+ under largely physiological conditions13,16,17. Here, we describe a protocol to study light stimulus-evoked cone Ca2+ dynamics in a transgenic Ca2+ biosensor mouse line (HR2.1:TN-XL), which expresses the FRET-based Ca2+ biosensor TN-XL18 selectively in cones, under the human red opsin promoter HR2.119. To access the cone terminals, an ex-vivo slice preparation20 was employed. The protocol was already successfully used in three studies on cone function in healthy mice10,21,22. Moreover, the protocol allows studying cone Ca2+ signaling in specific genetic conditions, e.g., by crossbreeding mouse models for hereditary retinal degeneration with HR2.1:TN-XL mice.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>High vacuum grease</td>
			<td>Dow Corning</td>
			<td>1018817</td>
			<td><a href="http://www.dowcorning.com/">http://www.dowcorning.com</a></td>
		</tr>
		<tr>
			<td>Cover slips</td>
			<td>R. Langenbrinck</td>
			<td></td>
			<td>24 x 60 mm; http://www.langenbrinck.com</td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>R. Langenbrinck</td>
			<td></td>
			<td>76 x 26 mm; http://www.langenbrinck.com</td>
		</tr>
		<tr>
			<td>Blades for tissue chopper</td>
			<td>MARTOR KG</td>
			<td>ARGENTAX No. 1044</td>
			<td>0.25 mm; http://www.martor.de</td>
		</tr>
		<tr>
			<td>Tissue chopper</td>
			<td>Custom-build</td>
			<td></td>
			<td>Based on a design by F. Werblin23, adapted by T. Schubert</td>
		</tr>
		<tr>
			<td>Nitrocellulose filter membrane gridded</td>
			<td>Merck Millipore</td>
			<td>AABG01300</td>
			<td>Filter type: 0.8 &#181;m; http://www.emdmillipore.com</td>
		</tr>
		<tr>
			<td>Isoflurane CP</td>
			<td>CP pharma</td>
			<td>AP/DRUGS/220/96</td>
			<td><a href="http://www.cp-pharma.de/">http://www.cp-pharma.de</a></td>
		</tr>
		<tr>
			<td>UV/green LED-based full-field stimulator</td>
			<td>Custom-build</td>
			<td></td>
			<td>for details, see10</td>
		</tr>
		<tr>
			<td>Open source microprocessor board</td>
			<td>Arduino</td>
			<td></td>
			<td><a href="http://www.arduino.cc/">http://www.arduino.cc</a></td>
		</tr>
		<tr>
			<td>Imaging software CfNT</td>
			<td>Custom-written</td>
			<td></td>
			<td>developed by Michael M&#252;ller, MPI for Medical Research, Heidelberg, Germany</td>
		</tr>
		<tr>
			<td>IgorPro</td>
			<td>Wavemetrics</td>
			<td></td>
			<td><a href="http://www.wavemetrics.com/">http://www.wavemetrics.com</a></td>
		</tr>
		<tr>
			<td>VC3-4 System focal perfusion system</td>
			<td>ALA Scientific Instrument</td>
			<td></td>
			<td>with 100 &#956;m-diameter tip manifold; http://www.alascience.com/</td>
		</tr>
		<tr>
			<td>Mai Tai HP DeepSee (Ti:Sapphire laser)</td>
			<td>Newport Spectra-Physics</td>
			<td></td>
			<td><a href="http://www.spectra-physics.com/">http://www.spectra-physics.com</a></td>
		</tr>
		<tr>
			<td>Movable objective microscope (MOM), 
			Designed by W. Denk, MPImF, Heidelberg, Germany</td>
			<td>Sutter Instruments</td>
			<td></td>
			<td><a href="http://www.sutter.com/MICROSCOPES/index.html">http://www.sutter.com/MICROSCOPES/index.html</a></td>
		</tr>
		<tr>
			<td>XLUMPlanFL 20x/0.95w objective</td>
			<td>Olympus</td>
			<td></td>
			<td><a href="http://www.olympusamerica.com/">http://www.olympusamerica.com</a></td>
		</tr>
		<tr>
			<td>Vaporizer 100 series</td>
			<td>Surgivet</td>
			<td></td>
			<td><a href="http://www.surgivet.com/">http://www.surgivet.com</a></td>
		</tr>
		<tr>
			<td>Ionomycin, Calcium salt</td>
			<td>Life technologies</td>
			<td>I24222</td>
			<td><a href="http://www.lifetechnologies.com/">http://www.lifetechnologies.com</a></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma-Aldrich</td>
			<td>E3889</td>
			<td><a href="https://www.sigmaaldrich.com/">https://www.sigmaaldrich.com</a></td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma-Aldrich</td>
			<td>101191250</td>
			<td><a href="http://www.sigmaaldrich.com/">http://www.sigmaaldrich.com</a></td>
		</tr>
		<tr>
			<td>Leica MZ95</td>
			<td>leica microsystems</td>
			<td></td>
			<td><a href="http://www.leica-microsystems.com/">http://www.leica-microsystems.com/</a></td>
		</tr>
	</tbody>
</table>

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.

By combining vertical slices (Figure 1) prepared from HR2.1:TN-XL mouse retina with two-photon imaging (Figure 2), we were able to perform ratiometric measurements of light stimulus-evoked Ca2+ signals in cone photoreceptor axon terminals. This approach represents a significant improvement in accessing and visualizing mouse cones over the pre-existing optical imaging methods with synthetic Ca2+ indicators. For instance, the cone-specific expression of the genetically-encoded ratiometric Ca2+ biosensor TN-XL greatly facilitated the identification of cone axon terminals (see ROIs in Figure 3A). Therefore, our protocol eliminates the risk of recording signals of unclear origin, as for example &#8220;mixed&#8221; signals with contributions from both cone and rod axon terminals.
Our approach allows resolving single-trial responses at the level of the individual cone axon terminal. Light-evoked extrusion of Ca2+ from the cone terminal is marked by an increase and a decrease in donor and acceptor fluorescence, respectively (Figure 3B).&#160;Decrease in fluorescence ratio (R) corresponds to the extrusion of Ca2+ from the cone terminal (Figure 3C), caused by the light-induced hyperpolarization of the cone and subsequent closure of voltage-gated Ca2+ channels. Because several cone axon terminals can be monitored simultaneously (Figure 3A), data acquisition is very efficient and hundreds of cone Ca2+ responses can be collected in a single experiment. By assessing these responses quantitatively (Figure 4), cone Ca2+ signals can be compared and evaluated in a range of conditions (see Discussion).
It is often sufficient to use the ratio between FRET-acceptor (citrine) and -donor (eCFP) fluorescence (FA/FD; for details, see 3.4) as a relative measure of the intracellular Ca2+ level. Yet, whenever necessary, the ratio can be calibrated to obtain absolute intracellular Ca2+ concentrations ([Ca2+]) (Figure 5). At the background illumination used in this protocol (for details, see10 and Figure&#160;3 legend), calibration yielded an estimate for the absolute resting [Ca2+] in &#8220;wild-type&#8221; HR2.1:TN-XL mouse cone terminals of 243 &#177; 159 nM (mean &#177; S.D.), which is within the range reported for mice in the literature11.
<img alt="Figure 1" src="/files/ftp_upload/52588/52588fig1.jpg" /> 
Figure 1.&#160;Preparation of vertical retinal slices. (A) Isolated and flattened retina prepared for slicing. (B) Rectangular piece of retina (see red box in A) mounted on filter paper membrane (dark grey) after slicing. (C) Top view of the membrane-mounted retinal slice fixed on a glass coverslip using grease. (D) Schematic drawing of a part of a retinal slice (see red box in C). Cone photoreceptors are indicated in green. V, ventral; D, dorsal; N, nasal; T, temporal; OS, outer segment; ONL, outer nuclear layer; OPL, outer plexiform layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer. <a href="https://www.jove.com/files/ftp_upload/52588/52588fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52588/52588fig2.jpg" /> 
Figure 2.&#160;Two-photon imaging of retinal slices: Illustration of the recording configuration. Top: Water-immersion objective lens focusing the beam of the scanning laser (red downward arrow) into the retina, and collecting the emitted fluorescence (upward arrows). The lens also collects transmitted infrared light from an illumination LED mounted below the condenser when imaging the slice using a CCD camera. Center: Retinal slice mounted in the recording chamber and perfused with extracellular solution. Bottom: Condenser lens focusing the light from the stimulation LEDs (and the infrared LED for CCD camera imaging) through the transparent bottom of the recording chamber into the retinal tissue. IR LED, infrared light-emitting diode; CCD, charge-coupled device; BP, band-pass. <a href="https://www.jove.com/files/ftp_upload/52588/52588fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52588/52588fig3.jpg" /> 
Figure 3.&#160;Light-evoked Ca2+ responses in axon terminals of mouse cone photoreceptors. (A) Left: Recordings focused on synaptic terminals (red box) of cone photoreceptors (green) in retinal slices. Right: Example for a recorded region with 7 individual terminals. Fluorescence was recorded using two channels: one for the FRET-acceptor citrine (top; 535 BP 50) and one for the FRET-donor eCFP (bottom; 480 BP 32). (B) Light-evoked changes in citrine (FA) and eCFP (FD) fluorescence recorded in a single ROI.&#160;(C) Ca2+ responses (as ratio FA/FD) of a cone terminal to a series of 1 sec bright light flashes in 5 sec intervals. ROI, region of interest; BP, band-pass filter; F, fluorescence intensity; a.u., arbitrary units; FRET, F&#246;rster resonance energy transfer; eCFP, enhanced cyan fluorescence protein. Stimulus&#160;intensity (as photo-isomerization rate in 103&#183;P&#183;s-1 per cone): 13.0 and 12.8 for M- and S-opsins, respectively, adding onto a background level (= LEDs off, excitation laser scanning) of ~10. <a href="https://www.jove.com/files/ftp_upload/52588/52588fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52588/52588fig4.jpg" /> 
Figure 4. Exemplary quantitative analysis of light-evoked cone Ca2+ responses. (A) Light-evoked Ca2+ responses (as &#916;R) from a single cone terminal (single trials: grey traces, n=16; mean: black trace). (B) Parameters determined for the mean Ca2+ response: Resting Ca2+ level (Rbase) representing the baseline prior to light stimulation, response size as area-under-the-curve (RA) and peak amplitude (Ramp), kinetics of response onset (trise, time interval from 20% to 80% of Ramp) and offset (tdecay, time interval from 80% to 20% of Ramp). <a href="https://www.jove.com/files/ftp_upload/52588/52588fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/52588/52588fig5.jpg" /> 
Figure 5. Estimating absolute Ca2+ concentration. Resting Ca2+ concentration ([Ca2+], as ratio FA/FD) recorded in 8 cone axon terminals in a TN-XL mouse (circles, with mean &#177; S.D.) for different extracellular solutions: twice in standard extracellular medium (Ctrl 1 and 2), then with minimal (&#8220;zero&#8221;) [Ca2+] (10 mM EGTA) and with high [Ca2+] (2.5 mM), the latter two conditions in the presence of 5 &#181;M ionomycin to equilibrate extracellular and intracellular Ca2+. <a href="https://www.jove.com/files/ftp_upload/52588/52588fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Imaging Ca2+ Dynamics in Cone Photoreceptor Axon Terminals of the Mouse Retina at JoVE.com

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.

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 ...

imaging-ca2-dynamics-cone-photoreceptor-axon-terminals-mouse

Research

JoVE Journal

Neuroscience

10.6K vues.  University of Tübingen.  Nous décrivons un protocole à suivre Ca 2+ dynamique dans les terminaisons axonales des cônes photorécepteurs utilisant une préparation de tranches ex-vivo de la rétine de souris. Ce protocole permet à des études approfondies de cône Ca 2+ signalisation dans un important système modèle mammifère, la souris. 

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

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

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 ...

Transretinal ERG Recordings from Mouse Retina: Rod and Cone Photoresponses

There are two distinct classes of image-forming photoreceptors in the vertebrate retina: rods and cones. Rods are able to detect single photons of light whereas cones operate continuously under rapidly changing bright light conditions. Absorption of light by rod- and cone-specific visual pigments in the outer segments of photoreceptors triggers a phototransduction cascade that eventually leads to closure of cyclic nucleotide-gated channels on the plasma membrane and cell hyperpolarization. This light-induced change in membrane current and potential can be registered as a photoresponse, by either classical suction electrode recording technique1,2 or by transretinal electroretinogram recordings (ERG) from isolated retinas with pharmacologically blocked postsynaptic response components3-5. The latter method allows drug-accessible long-lasting recordings from mouse photoreceptors and is particularly useful for obtaining stable photoresponses from the scarce and fragile mouse cones. In the case of cones, such experiments can be performed both in dark-adapted conditions and following intense illumination that bleaches essentially all visual pigment, to monitor the process of cone photosensitivity recovery during dark adaptation6,7. In this video, we will show how to perform rod- and M/L-cone-driven transretinal recordings from dark-adapted mouse retina. Rod recordings will be carried out using retina of wild type (C57Bl/6) mice. For simplicity, cone recordings will be obtained from genetically modified rod transducin &alpha;-subunit knockout (T&alpha;-/-) mice which lack rod signaling8.

We describe a relatively simple method of transretinal electroretinogram (ERG) recordings for obtaining rod and cone photoresponses from intact mouse retina. This approach takes advantage of the block of synaptic transmission from photoreceptors to isolate their light responses and record them using field electrodes placed across the isolated flat-mounted retina.

There are two distinct classes of image-forming photoreceptors in the vertebrate retina: rods and cones. Rods are able to detect single photons of light ...

Use of pHluorin to Assess the Dynamics of Axon Guidance Receptors in Cell Culture and in the Chick Embryo

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation of the guidance receptors is the first step of the signaling mechanisms allowing axon tips, the growth cones, to respond to the ligands. As such, the modulation of their availability at the cell surface is one of the mechanisms that participate in setting the growth cone sensitivity. We describe here a method to precisely visualize the spatio-temporal cell surface dynamics of an axon guidance receptor both in vitro and in vivo in the developing chick spinal cord. We took advantage of the pH-dependent fluorescence property of a green fluorescent protein (GFP) variant to specifically detect the fraction of the axon guidance receptor that is addressed to the plasma membrane. We first describe the in vitro validation of such pH-dependent constructs and we further detail their use in vivo, in the chick spinal chord, to assess the spatio-temporal dynamics of the axon guidance receptor of interest.

We describe here the use of a pH-sensitive green fluorescent protein variant, pHluorin, to study the spatio-temporal dynamics of axon guidance receptors trafficking at the cell surface.&#160;The pHluorin-tagged receptor is expressed both in cell culture and in vivo, using electroporation of the chick embryo.

During development, axon guidance receptors play a crucial role in regulating axons sensitivity to both attractive and repulsive cues. Indeed, activation ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...

Vibratome Sectioning Mouse Retina to Prepare Photoreceptor Cultures

The retina is a part of the central nervous system that has organized architecture, with neurons in layers from the photoreceptors, both rods and cones in contact with the retinal pigmented epithelium in the most distant part on the retina considering the direction of light, and the ganglion cells in the most proximal distance. This architecture allows the isolation of the photoreceptor layer by vibratome sectioning. The dissected neural retina of a mouse aged 8 days is flat-embedded in 4% gelatin on top of a slice of 20% gelatin photoreceptor layer facing down. Using a vibratome and a double edged razor blade, the 100 &#181;m thick inner retina is sectioned. This section contains the ganglion cells and the inner layer with notably the bipolar cells. An intermediary section of 15 &#181;m is discarded before 200 &#181;m of the outer retina containing the photoreceptors is recovered. The gelatin is removed by heating at 37 &#176;C. Pieces of outer layer are incubated in 500 &#181;l of Ringer&#39;s solution with 2 units of activated papain for 20 min at 37 &#176;C. The reaction is stopped by adding 500 &#181;l 10% fetal calf serum (FCS) in Dulbecco&#39;s Modified Eagle Medium (DMEM), then 25 units of DNAse I is added before centrifugation at RT, washed several times to remove serum and the cells are resuspended in 500 &#181;l of DMEM and seeded at 1 x 105 cells/cm2. The cells are grown to 5 days in vitro and their viability scored using live/dead assay. The purity of the culture is first determined by microscopic observation during the experiment. The purity is then validated by seeding and fixing cells on a histological slide and analyzing using a rabbit polyclonal anti-SAG, a photoreceptor marker and mouse monoclonal anti-RHO, a rod photoreceptor specific marker. Alternatively, the photoreceptor layer (97% rods) can be used for gene or protein expression analysis and for transplantation.

Neural retina of a mouse aged 8 days is on top of a 4% gelatin block. After isolation of the photoreceptor layer (200 &#181;m) by vibratome, the photoreceptors are seeded after mechanical and enzymatic dissociation for culture. The photoreceptor layer can be used for molecular, biochemical analyses or transplantation.

The retina is a part of the central nervous system that has organized architecture, with neurons in layers from the photoreceptors, both rods and cones in ...

Whole-mount Imaging of Mouse Embryo Sensory Axon Projections

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we describe a method to label in situ a subset of dorsal root ganglion (DRG) axon projections to assess their phenotypic characteristics using several genetically manipulated mouse lines. The TrkA-positive neurons are nociceptor neurons, dedicated to the transmission of pain signals. We utilize a TrkAtaulacZ mouse line to label the trajectories of all TrkA-positive peripheral axons in the intact mouse embryo. We further breed the TrkAtaulacZ line onto a Bax null background, which essentially abolishes neuronal apoptosis, in order to assess growth-related questions independently of possible effects of genetic manipulations on neuronal survival. Subsequently, genetically modified mice of interest are bred with the TrkAtaulacZ/Bax null line and are then ready for study using the techniques described herein. This presentation includes detailed information on mouse breeding plans, genotyping at the time of dissection, tissue preparation, staining and clearing to allow for visualization of full-length axonal trajectories in whole-mount preparation.

We present here an optimized protocol to genotype, stain and prepare fetal mice for the imaging of peripheral nociceptor axon projections in the whole animal, as an effective method to assess sensory axon growth phenotypes in developing genetically engineered mice.

The visualization of full-length neuronal projections in embryos is essential to gain an understanding of how mammalian neuronal networks develop. Here we ...

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 ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...