Name: cut-loading: a useful tool for examining the extent of gap junction tracer coupling between retinal neurons
Uploaded: 2012-01-12T12:00:00
Duration: 10 min 11 s
Description: Watch this Scientific Journal Video about Cut-loading: A Useful Tool for Examining the Extent of Gap Junction Tracer Coupling Between Retinal Neurons at JoVE.com

This work was funded by NIH grant EY005102 to S.C.M. and EY018640 to C.P.R.

1. Intact neural retina preparations for goldfish, mice and rabbits
<ol>
	<li>Perform all of the following steps, including those described in "2) Cut-loading," under constant ambient (background) illumination. Please note that for the purposes of this protocol, the procedures are described as performed in constant darkness (i.e. under very dark (i.e. "scotopic") conditions &le; 0.0001 lux) using night vision infrared goggles, but other higher intensity levels of constant ambient illumination can be used instead to determine the effect of other levels of ambient illumination on gap junction tracer coupling.</li>
	<li>Dark-adapt the experimental animal (goldfish, mouse or rabbit) for at least 1 hr before surgery.</li>
	<li>As described previously7, deeply anesthetize goldfish by placing them in tricaine methanesulfonate (MS222; 150 mg/L of buffered (sodium bicarbonate-containing) fish tank water), and deeply anesthetize mice with ketamine (100 mg/kg, i.p.) and rompun (10 mg/kg, i.p.). Deeply anesthetize rabbits with urethane (loading dose: 2.0 g/kg, i.p.) and also use local intraorbital anesthesia (2% Xylocaine), as described previously8. All experimental procedures involving the care and use of animals should be performed in accordance with federal guidelines and be reviewed and approved by local university animal care and use committees. (Note: In the experiments described here, all experimental procedures involving the care and use of fish, mice and rabbits were performed in accordance with NIH guidelines and were reviewed and approved by Ohio State University Institutional Animal Care and Use Committee.)</li>
	<li>For fish, mice and rabbits, following enucleation, remove the anterior portion of each eyeball. Then, place a piece of filter paper on top of the posterior portion of the eye and invert the filter paper and eye, so that the filter paper is underneath the posterior portion of the eye. Then, using fine forceps dissect the intact neural retina from the posterior portion of the eye by gently peeling away the pigment epithelium/choroid/sclera, which are attached to each other, from the neural retina, which is attached to the filter paper. As this is done, cut the optic nerve using fine spring-loaded scissors. The neural retina, oriented photoreceptor-side up, should now be attached to the filter paper and separated from the rest of the eye.</li>
</ol>
2. Cut-loading
<ol>
	<li>Submerge the retinas in oxygenated Ringer's solution (Table 1) in a 6-well plate (~5 mL/well) for 30 min under constant ambient (background) illumination (e.g. under very dark (i.e. "scotopic") conditions) with or without a test drug. Maintain fish retinas in oxygenated (5% CO2 /95% O2) bicarbonate-based Ringer's at 22&deg;C by sealing the 6-well plate with parafilm and maintain mouse and rabbit retinas at 36&deg;C in a 5% CO2 /95% O2 incubator. When a test drug is used, it should be present during all subsequent steps until fixation</li>
	<li>Prepare the tracer solution (typically 100 &mu;L) by dissolving neurobiotin (0.5%), a biotinylated molecule, in Ringer's solution immediately before cutting through the retina. Note that 0.5% rhodamine dextran (high (&gt; 10,000) MW) may be added to the solution. Due to its high molecular weight, rhodamine dextran does not cross gap junctions and only labels cells that have been damaged by the cut. As shown in Fig. 3, this is a useful way to distinguish cells that have accumulated neurobiotin because they have been injured, from those that have accumulated the tracer by diffusion through gap junctions.</li>
	<li>Place the neurobiotin solution on a glass (or plastic) Petri dish, tap the filter paper, to which the retina is attached, on a paper towel to remove excess Ringer, dip a razor blade (or other sharp blade) in the neurobiotin solution and then make a radial cut through the retina and filter paper. If preferred, spacers can be used so that the cut is made through the retina, but not the filter paper. The cut should be oriented perpendicular to the retinal surface. Dip the razor blade in the neurobiotin solution again and cut the retina a second time. Repeat this up to 4 cuts per retina (see Fig. 1). Use a dissecting microscope when making razor cuts through mouse and other small retinas.</li>
	<li>As shown in Fig. 1, submerge the retina again in fresh Ringer's medium, with or without test drug, using a 6-plate well (5 mL Ringer/well). Incubate the retina for 15 min to allow for loading and diffusion.</li>
	<li>Wash three times for 5 min each with fresh Ringer's medium with or without test drug.</li>
	<li>Fix the retina with 4% paraformaldehyde in 0.1 M phosphate buffer (PBS, pH 7.4) for 1 hr at RT.</li>
	<li>Wash with 0.1 M PBS overnight at 4&deg;C.</li>
	<li>The following day, react with 2% streptavidin-Alexa 488 (final concentration 10 &mu;g/mL) in 0.1M PBS + 0.3% Triton X-100, and keep overnight at 4&deg;C.</li>
	<li>Wash three times for 10 min each with 0.1M PBS at RT</li>
	<li>In PBS, detach the retina from the filter paper and then, using a fine brush, place the retina, oriented photoreceptor-side up, onto a slide.</li>
	<li>Gently mount with Vectashield mounting medium (Vector Laboratories, Burlingame, CA).</li>
	<li>Take pictures with a laser scanning confocal microscope (e.g. Zeiss 510 META) at the same magnification, resolution and settings for every experimental condition, so that you can compare the effects of different experimental conditions (e.g. test drugs; illumination conditions) on the extent of gap junction tracer coupling (Fig. 2A-C and Fig. 4A-C). Although a single image may contain all of the labeled cells, it is recommended that you collect a z-stack of the area of interest and collapse it into a single image.</li>
</ol>
3. Quantification of tracer coupling using ImageJ
<ol>
	<li>In the LSM image browser, open the image, click EXPORT and save the picture as a TIF-16 bit NOT compressed file. Scale bar should be included in this TIF image.</li>
	<li>Using NIH ImageJ software, measure fluorescence intensity of Alexa-488-labeled neurobiotin from low-magnification images of whole-mount retinas. Open the TIF file using ImageJ software and then draw a straight line corresponding to the scale bar. Go to ANALYSE, SET SCALE and enter the known distance and unit of length, then click OK. Now measurement results can be presented in calibrated units, such as millimeters.</li>
	<li>Select the area of interest using the rectangular selection tool. The peak of fluorescence should be positioned at the left border of your selection. Make sure that there is no fluorescence signal near the right border. If not, rotate the active image (IMAGE, then ROTATE).</li>
	<li>Click ANALYSE and PLOT PROFILE. You should see a curve with an exponential decay from the left to the right.</li>
	<li>Click COPY in the pop-up window and paste it in EXCEL. Now you see two columns showing the distance from the cut (left column) and the fluorescence intensity as a function of the distance from the cut (right column).</li>
	<li>Divide each raw fluorescence value by the maximum fluorescence value to obtain the relative fluorescence intensity.</li>
	<li>Copy two columns corresponding to the distance from the cut (X) and the relative fluorescence intensity (Y), and then paste them into Origin software.</li>
	<li>Fit with the EXPONENTIAL DECAY #1 function (Fig. 2D and Fig. 4D), which is in the form: 
	Y = Y0 + Ymax*exp(-x/&lambda;) 
	where Y is the relative fluorescence intensity, Yo is the background fluorescence, Ymax is the maximum relative fluorescence, &lambda; is the space (length) constant, and x is the distance from the cut.</li>
	<li>Compare the space constant (&lambda;) values at different experimental conditions using the t-test or ANOVA (Fig. 2E and Fig. 4E). Note that alternative techniques of quantification have been described by others13,14.</li>
</ol>
4. Representative Results
Representative examples of photoreceptor cell tracer coupling as determined by cut-loading are presented in Figures 2 and 3 (fish) and Figure 4 (rabbit). Confocal images were taken using the same settings for comparison (Fig. 2A-C and Fig. 4A-C) and the fluorescence intensity was plotted as a function of the distance from the cut and fitted by the exponential function shown in No. 3.8 above (Fig. 2D and Fig. 4D). Space constant values for each condition are shown in Figures 2E and 4E, illustrating that the extent of gap junction tracer coupling can be quantified using the cut-loading technique. In addition, the results are highly reproducible. Use of the cut-loading technique as a means of quantifying the extent of gap junction tracer coupling is also validated by the finding that fluorescence intensity decreases exponentially as a function of distance from the cut in all cases examined7,8 (see also Figs. 2D and 4D here), indicating that neurobiotin entered the photoreceptors via the razor cut and not from other retinal sites. Moreover, the qualitatively similar day/night difference in photoreceptor cell tracer coupling observed in goldfish with tracer injections into single cones and with cut-loading7 substantiates cut-loading as a relatively accurate means of measuring the extent of photoreceptor coupling.
The cut-loading technique can also be used to investigate other types of electrical synapses in the retina. For example, Figure 5 illustrates that rabbit A-type (Fig. 5A) and B-type (Fig. 5B) horizontal cells exhibit homologous tracer coupling following cut-loading and diffusion of neurobiotin under dark-adapted conditions.
<table border="1">
	<tbody>
		<tr>
			<td>Compound</td>
			<td>Fish</td>
			<td>Mouse</td>
			<td>Rabbit</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>130</td>
			<td>120</td>
			<td>117</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>20</td>
			<td>25</td>
			<td>30</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>-</td>
			<td>1</td>
			<td>0.5</td>
		</tr>
		<tr>
			<td>KCL</td>
			<td>2.5</td>
			<td>5</td>
			<td>3.1</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>10</td>
			<td>10</td>
			<td>10</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>MgSO4-7H2O</td>
			<td>-</td>
			<td>1</td>
			<td>1.2</td>
		</tr>
		<tr>
			<td>Glutamine</td>
			<td>-</td>
			<td>0.1</td>
			<td>0.1</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>0.7</td>
			<td>2</td>
			<td>2</td>
		</tr>
	</tbody>
</table>
Table 1: Composition of the Ringer's solutions for goldfish, mouse and rabbit retinas. The concentrations are presented in mM. The Ringer's solutions are bubbled with 5% CO2 /95% O2 and maintained at 22 &deg;C (fish) or 36 &deg;C (mammals). "-" : not included in the Ringer for this species.
<img alt="Figure 1" src="/files/ftp_upload/3180/3180fig1.jpg" />
Figure 1.: Flow chart showing the cut-loading procedure. After isolation of the intact neural retina, several radial cuts were made by a blade that was first dipped in 0.5% neurobiotin solution. The retina was incubated for tracer loading and diffusion, and then washed before fixation with 4% paraformaldehyde (PFA) in 0.1M phosphate buffer (PB). Tracer coupling was examined using streptavidin-conjugated Alexa-488.
<img alt="Figure 2" src="/files/ftp_upload/3180/3180fig2.jpg" />
Figure 2. Day-night difference in photoreceptor tracer coupling in goldfish is revealed by the cut-loading technique. Photoreceptor cell gap junction neurobiotin tracer coupling was extensive at night (B) and in the day following application of spiperone (10 &mu;M), a selective dopamine D2 receptor antagonist (C), but not in the day under control conditions (A). D) Normalized relative fluorescent intensity as a function of the distance from the cuts (indicated by arrows in A-C). E) Space constant values obtained from the data in D and other experiments (n = 4). ***P &lt; 0.001.
<img alt="Figure 3" src="/files/ftp_upload/3180/3180fig3.jpg" />
Figure 3. A representative example showing fluorescence in the photoreceptor cell layer of a dark-adapted goldfish retina at night following cut-loading with a solution of both neurobiotin and rhodamine dextran. Rhodamine dextran (shown in red), which does not diffuse through open gap junction channels due to its high molecular weight (&gt; 10,000 MW), only labeled cells near the cut. In contrast, neurobiotin (shown in green) diffused through gap junctions and can be seen in photoreceptor cells far from the cut. The location of the cut is indicated by the arrow in each panel. Scale bar: 200 &mu;m.
<img alt="Figure 4" src="/files/ftp_upload/3180/3180fig4.jpg" />
Figure 4. Day-night difference in photoreceptor tracer coupling in rabbit retina is revealed by the cut-loading technique. Photoreceptor cell gap junction neurobiotin tracer coupling was extensive at night (B) and in the day following application of spiperone (10 &mu;M) (C), but not in the day under control conditions (A). In A-C, perpendicular views of the 3D reconstruction of rabbit photoreceptors near the cut are shown. D) Normalized relative fluorescent intensity as a function of the distance from the cuts. E) Space constant values obtained from the data in D and other experiments (n = 3). ***P &lt; 0.001.
<img alt="Figure 5" src="/files/ftp_upload/3180/3180fig5.jpg" />
Figure 5. Cut-loading reveals that dark-adapted rabbit horizontal cells are tracer coupled. Both A-type (A) and B-type (B) horizontal cells in dark-adapted rabbit retinas exhibited homologous neurobiotin tracer coupling.

<ol>
	<li>Bennett, M. V. L., Zukin, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrical+coupling+and+neuronal+synchronization+in+the+mammalian+brain.">Electrical coupling and neuronal synchronization in the mammalian brain.</a> Neuron. 41, 495-511 (2004).</li><li>Connors, B. W., Long, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrical+synapses+in+the+mammalian+brain.">Electrical synapses in the mammalian brain.</a> Annu. Rev. Neurosci. 27, 393-418 (2004).</li><li>Vaney, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Many+diverse+types+of+retinal+neurons+show+tracer+coupling+when+injected+with+biocytin+or+Neurobiotin.">Many diverse types of retinal neurons show tracer coupling when injected with biocytin or Neurobiotin.</a> Neurosci. Lett. 125, 187-190 (1991).</li><li>Bloomfield, S. A., V&#246;lgyi, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+diverse+functional+roles+and+regulation+of+neuronal+gap+junctions+in+the+retina.">The diverse functional roles and regulation of neuronal gap junctions in the retina.</a> Nat. Rev. Neurosci. 10, 495-506 (2009).</li><li>Weiler, R., Pottek, M., He, S., Vaney, D. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+coupling+between+retinal+horizontal+cells+by+retinoic+acid+and+endogenous+dopamine.">Modulation of coupling between retinal horizontal cells by retinoic acid and endogenous dopamine.</a> Brain. Res. Brain. Res. Rev. 32, 121-129 (2000).</li><li>Xin, D., Bloomfield, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+nitric+oxide+on+horizontal+cells+in+the+rabbit+retina.">Effects of nitric oxide on horizontal cells in the rabbit retina.</a> Vis. Neurosci. 17, 799-811 (2000).</li><li>Ribelayga, C., Cao, Y., Mangel, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+circadian+clock+in+the+retina+controls+rod-cone+coupling.">The circadian clock in the retina controls rod-cone coupling.</a> Neuron. 59, 790-801 (2008).</li><li>Ribelayga, C., Mangel, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+a+circadian+clock-controlled+neural+pathway+in+the+rabbit+retina.">Identification of a circadian clock-controlled neural pathway in the rabbit retina.</a> PLoS One. 5, e11020-e11020 (2010).</li><li>Ouyang, X., Winbow, V. M., Patel, L. S., Burr, G. S., Mitchell, C. K., O'Brien, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+Kinase+A+mediates+regulation+of+gap+junctions+containing+connexin35+through+a+complex+pathway.">Protein Kinase A mediates regulation of gap junctions containing connexin35 through a complex pathway.</a> Brain. Res. Mol. Brain. Res. 135, 1-11 (2005).</li><li>Patel, L. S., Mitchell, C. K., Dubinsky, W. P., O'Brien, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+gap+junction+coupling+through+the+neuronal+connexin+Cx35+by+nitric+oxide+and+cGMP.">Regulation of gap junction coupling through the neuronal connexin Cx35 by nitric oxide and cGMP.</a> Cell. Commun. Adhes. 13, 41-54 (2006).</li><li>Peters, J. L., Cassone, V. M., Zoran, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Melatonin+modulates+intercellular+communication+among+cultured+chick+astrocytes.">Melatonin modulates intercellular communication among cultured chick astrocytes.</a> Brain. Res. 1031, 10-19 (2005).</li><li>Cusato, K., Bosco, A., Rozental, R., Guimar&#227;es, C. A., Reese, B. E., Linden, R., Spray, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gap+junctions+mediate+bystander+cell+death+in+developing+retina.">Gap junctions mediate bystander cell death in developing retina.</a> J. Neurosci. 23, 6413-6422 (2003).</li><li>Li, H., Chuang, A. Z., O'Brien, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoreceptor+coupling+is+controlled+by+connexin+35+phosphorylation+in+zebrafish+retina.">Photoreceptor coupling is controlled by connexin 35 phosphorylation in zebrafish retina.</a> J. Neurosci. 29, 15178-15186 (2009).</li><li>Mills, S. L., Massey, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+kinetics+of+tracer+movement+through+homologous+gap+junctions+in+the+rabbit+retina.">The kinetics of tracer movement through homologous gap junctions in the rabbit retina.</a> Vis. Neurosci. 15, 765-777 (1998).</li><li>Mills, S. L., Massey, S. 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No conflicts of interest declared.

The cut-loading method described here is a useful and straightforward technique to determine the extent of gap junction tracer coupling between retinal neurons under different illumination conditions and at different times of the day and night. Advantages of this technique include the ability to quantify the extent of gap junction tracer coupling between neurons in intact retinal tissue under a variety of illumination conditions during the day and night and to do so for coupled neurons that have small diameter somata. Limitations of the technique in the study of gap junctions in intact tissue fall into two general categories. First, tracer diffusion through open gap junctions may be relatively difficult to observe due to a) the small diameter of or the charge associated with the open channels and b) the relative volumes of coupled cellular compartments1,14,15. That is, tracer diffusion from a small cell to a larger cell or group of coupled cells, compared to tracer diffusion from a larger cell to a smaller cell, may be more difficult to detect due to tracer dilution. Second, under some physiological conditions, the extent of tracer diffusion through gap junctions in intact tissue may not accurately reflect the strength of gap junctional conductance due to differences in the conductance of small electric current-carrying ions, compared to the permeability of relatively large tracer molecules1,14,15. In general, evidence of tracer coupling strongly suggests the presence of functioning, open gap junction channels, but under some physiological conditions, tracer diffusion may not occur or be observed even though electrophysiological recordings suggest the presence of functioning, open gap junctions.
It seems likely that the cut-loading technique can also be used to investigate the extent of gap junction tracer coupling between neurons in intact tissue from other regions of the central nervous system.

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments (optional)</td>
		</tr>
		<tr>
			<td>Tricaine methane sulfonate (MS222)</td>
			<td>Sigma-Aldrich</td>
			<td>A5040</td>
			<td>150 mg/L of buffered fish tank water</td>
		</tr>
		<tr>
			<td>Urethane</td>
			<td>Sigma-Aldrich</td>
			<td>U2500</td>
			<td>2 g/kgloading dose</td>
		</tr>
		<tr>
			<td>Dual Tube Night Vision Goggle</td>
			<td>Night Optics USA</td>
			<td>D-221</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Filter Paper, Grade No. 4</td>
			<td>Whatman</td>
			<td>1004-090</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Fine Forceps, Dumont No. 5, Biologie, 11 cm long</td>
			<td>Fine Science Tools</td>
			<td>11295-10</td>
		</tr>
		<tr>
			<td>Fine Scissors, spring-loaded, 8 mm blade, straight</td>
			<td>Fine Science Tools</td>
			<td>15025-10</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Neurobiotin</td>
			<td>Vector</td>
			<td>SP-1120</td>
			<td>0.5%</td>
		</tr>
		<tr>
			<td>Streptavidin-conjugated Alexa 488</td>
			<td>Invitrogen</td>
			<td>S11223</td>
			<td>2%</td>
		</tr>
		<tr>
			<td>Dextran rhodamine(high (&gt; 10,000) MW)</td>
			<td>Invitrogen</td>
			<td>D1817</td>
			<td>0.5%</td>
		</tr>
		<tr>
			<td>Vectashield mounting medium</td>
			<td>Vector</td>
			<td>H-1000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Zeiss 510 META Laser Scanning Confocal Microscope</td>
			<td>Carl Zeiss, Inc.</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>LSM-5 Image Browser 3,2,0,115</td>
			<td>Carl Zeiss, Inc.</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>ImageJ Software</td>
			<td>NIH</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>OriginPro 8.0</td>
			<td>OriginLab Corp.</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>

cut-loading: a useful tool for examining the extent of gap junction tracer coupling between retinal neurons

In addition to chemical synaptic transmission, neurons that are connected by gap junctions can also communicate rapidly via electrical synaptic transmission. Increasing evidence indicates that gap junctions not only permit electrical current flow and synchronous activity between interconnected or coupled cells, but that the strength or effectiveness of electrical communication between coupled cells can be modulated to a great extent1,2. In addition, the large internal diameter (~1.2 nm) of many gap junction channels permits not only electric current flow, but also the diffusion of intracellular signaling molecules and small metabolites between interconnected cells, so that gap junctions may also mediate metabolic and chemical communication. The strength of gap junctional communication between neurons and its modulation by neurotransmitters and other factors can be studied by simultaneously electrically recording from coupled cells and by determining the extent of diffusion of tracer molecules, which are gap junction permeable, but not membrane permeable, following iontophoretic injection into single cells. However, these procedures can be extremely difficult to perform on neurons with small somata in intact neural tissue.
Numerous studies on electrical synapses and the modulation of electrical communication have been conducted in the vertebrate retina, since each of the five retinal neuron types is electrically connected by gap junctions3,4. Increasing evidence has shown that the circadian (24-hour) clock in the retina and changes in light stimulation regulate gap junction coupling3-8. For example, recent work has demonstrated that the retinal circadian clock decreases gap junction coupling between rod and cone photoreceptor cells during the day by increasing dopamine D2 receptor activation, and dramatically increases rod-cone coupling at night by reducing D2 receptor activation7,8. However, not only are these studies extremely difficult to perform on neurons with small somata in intact neural retinal tissue, but it can be difficult to adequately control the illumination conditions during the electrophysiological study of single retinal neurons to avoid light-induced changes in gap junction conductance.
Here, we present a straightforward method of determining the extent of gap junction tracer coupling between retinal neurons under different illumination conditions and at different times of the day and night. This cut-loading technique is a modification of scrape loading9-12, which is based on dye loading and diffusion through open gap junction channels. Scrape loading works well in cultured cells, but not in thick slices such as intact retinas. The cut-loading technique has been used to study photoreceptor coupling in intact fish and mammalian retinas7, 8,13, and can be used to study coupling between other retinal neurons, as described here.

Watch this Scientific Journal Video about Cut-loading: A Useful Tool for Examining the Extent of Gap Junction Tracer Coupling Between Retinal Neurons at JoVE.com

Cut-loading: A Useful Tool for Examining the Extent of Gap Junction Tracer Coupling Between Retinal Neurons

An easy and convenient method to determine the extent of gap junction tracer coupling between retinal neurons is described. This technique enables one to investigate the function of the electrical synapses between neurons in the intact retina under different illumination conditions and at different times of the day and night.

In addition to chemical synaptic transmission, neurons that are connected by gap junctions can also communicate rapidly via electrical synaptic ...

cut-loading-useful-tool-for-examining-extent-gap-junction-tracer

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Neuroscience

The Organotypic Hippocampal Slice Culture Model for Examining Neuronal Injury

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of the hippocampus is relatively preserved 1. The organotypic culture system allows for the examination of neuronal, astrocytic and microglial effects, but as an ex vivo preparation, does not address effects of blood flow, or recruitment of peripheral inflammatory cells. To that end, this culture method is frequently used to examine excitotoxic and hypoxic injury to pyramidal neurons of the hippocampus, but has also been used to examine the inflammatory response. Herein we describe the methods for generating hippocampal slice cultures from postnatal rodent brain, administering toxic stimuli to induce neuronal injury, and assaying and quantifying hippocampal neuronal death.

The organoptypic hippocampal slice culture model is an in vitro model used to examine neuronal injury in a variety of paradigms. In this article, we describe the methods for generating slice cultures and quantifying neuronal injury.

Organotypic hippocampal slice culture is an in vitro method to examine mechanisms of neuronal injury in which the basic architecture and composition of ...

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and cones). The RPE is a monolayer of pigmented cuboidal cells and associates closely with the neural retina just above it. This association makes the RPE of great interest to researchers studying retinal diseases. The RPE is also the site of an in vivo assay of homology-directed DNA repair, the pun assay. The mouse eye is particularly difficult to dissect due to its small size (about 3.5mm in diameter) and its spherical shape. This article demonstrates in detail a procedure for dissection of the eye resulting in a whole mount of the RPE. In this procedure, we show how to work with, rather than against, the spherical structure of the eye. Briefly, the connective tissue, muscle, and optic nerve are removed from the back of the eye. Then, the cornea and lens are removed. Next, strategic cuts are made that result in significant flattening of the remaining tissue. Finally, the neural retina is gently lifted off, revealing an intact RPE, which is still attached to the underlying choroid and sclera. This whole mount can be used to perform the pun assay or for immunohistochemistry or immunofluorescent assessment of the RPE tissue.

A formal demonstration of the dissection of a mouse eye, resulting in a whole mount of the retinal pigment epithelium.

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and ...

Single-cell Profiling of Developing and Mature Retinal Neurons

Highly specialized, but exceedingly small populations of cells play important roles in many tissues. The identification of cell-type specific markers and gene expression programs for extremely rare cell subsets has been a challenge using standard whole-tissue approaches. Gene expression profiling of individual cells allows for unprecedented access to cell types that comprise only a small percentage of the total tissue1-7. In addition, this technique can be used to examine the gene expression programs that are transiently expressed in small numbers of cells during dynamic developmental transitions8.
This issue of cellular diversity arises repeatedly in the central nervous system (CNS) where neuronal connections can occur between quite diverse cells9. The exact number of distinct cell types is not precisely known, but it has been estimated that there may be as many as 1000 different types in the cortex itself10. The function(s) of complex neural circuits may rely on some of the rare neuronal types and the genes they express. By identifying new markers and helping to molecularly classify different neurons, the single-cell approach is particularly useful in the analysis of cell types in the nervous system. It may also help to elucidate mechanisms of neural development by identifying differentially expressed genes and gene pathways during early stages of neuronal progenitor development.
As a simple, easily accessed tissue with considerable neuronal diversity, the vertebrate retina is an excellent model system for studying the processes of cellular development, neuronal differentiation and neuronal diversification. However, as in other parts of the CNS, this cellular diversity can present a problem for determining the genetic pathways that drive retinal progenitors to adopt a specific cell fate, especially given that rod photoreceptors make up the majority of the total retinal cell population11. Here we report a method for the identification of the transcripts expressed in single retinal cells (Figure 1). The single-cell profiling technique allows for the assessment of the amount of heterogeneity present within different cellular populations of the retina2,4,5,12. In addition, this method has revealed a host of new candidate genes that may play role(s) in the cell fate decision-making processes that occur in subsets of retinal progenitor cells8. With some simple adjustments to the protocol, this technique can be utilized for many different tissues and cell types.

A method for the isolation of single retinal cells and subsequent amplification of their cDNAs is described. Single-cell transcriptomics reveals the degree of cellular heterogeneity present in a tissue and uncovers new marker genes for rare cell populations. The accompanying protocol can be adjusted to suit many different cell types.

Highly specialized, but exceedingly small populations of cells play important roles in many tissues. The identification of cell-type specific markers and ...

Antibody Transfection into Neurons as a Tool to Study Disease Pathogenesis

Antibodies provide the ability to gain novel insight into various events taking place in living systems. The ability to produce highly specific antibodies to target proteins has allowed for very precise biological questions to be addressed. Importantly, antibodies have been implicated in the pathogenesis of a number of human diseases including systemic lupus erythematosus (SLE), rheumatoid arthritis (RA), paraneoplastic syndromes, multiple sclerosis (MS) and human T-lymphotropic virus type 1 (HTLV-1) associated myelopathy/tropical spastic paraparesis (HAM/TSP) 1-9. How antibodies cause disease is an area of ongoing investigation, and data suggests that interactions between antibodies and various intracellular molecules results in inflammation, altered cellular messaging, and apoptosis 10. It has been shown that patients with MS and HAM/TSP produce autoantibodies to the intracellular RNA binding protein heterogeneous ribonuclear protein A1 (hnRNP A1) 3, 5-7, 9, 11. Recent data indicate that antibodies to both intra-neuronal and surface antigens are pathogenic 3, 5-9, 11. Thus, a procedure that allows for the study of intracellular antibody:protein interactions would lend great insight into disease pathogenesis.
Genes are commonly transfected into primary cells and cell lines in culture, however transfection of antibodies into cells has been hindered by alteration of antibody structure or poor transfection efficiency 12. Other methods of transfection include antibody transfection based on cationic liposomes (consisting of DOTAP/DOPE) and polyethylenimines (PEI); both of which resulted in a ten-fold decrease in antibody transfection compared to controls 12. The method performed in our study is similar to cationic lipid-mediated methods and uses a lipid-based mechanism to form non-covalent complexes with the antibodies through electrostatic and hydrophobic interactions 13. We utilized Ab-DeliverIN reagent, which is a lipid formulation capable of capturing antibodies through non-covalent electrostatic and hydrophobic interactions and delivering them inside cells. Thus chemical and genetic couplings are not necessary for delivery of functional antibodies into living cells. This method has enabled us to perform various antibody tracing and protein localization experiments, as well as the analyses of the molecular consequences of intracellular antibody:protein interactions 9.
In this protocol, we will show how to transfect antibodies into neurons rapidly, reproducibly and with a high degree of transfection efficiency. As an example, we will use anti-hnRNP A1 and anti-IgG antibodies. For easy quantification of transfection efficiency we used anti-hnRNP A1 antibodies labelled with Atto-550-NHS and FITC-labeled IgG. Atto550 NHS is a new label with high molecular absorbtion and quantum yield. Excitation source and fluorescent filters for Atto550 are similar to Cy3 (Ex. 556 Em. 578). In addition, Atto550 has high photostability. FITC-labeled IgG were used as a control to show that this method is versatile and not dye dependent. This approach and the data that is generated will assist in understanding of the role that antibodies to intracellular target antigens might play in the pathogenesis of human diseases.

A rapid approach to investigate interactions and effects on molecular mechanisms related to the presence of antibodies in an intracellular environment is described. The method involves transfection of antibodies into live cells using a non-covalent complex formation based on a lipid formulation. The technique is adaptable to immortalized cell lines and primary cells.

Antibodies provide the ability to gain novel insight into various events taking place in living systems.  The ability to produce highly specific ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

A Dual Tracer PET-MRI Protocol for the Quantitative Measure of Regional Brain Energy Substrates Uptake in the Rat

We present a method for comparing the uptake of the brain&#39;s two key energy substrates: glucose and ketones (acetoacetate [AcAc] in this case) in the rat. The developed method is a small-animal positron emission tomography (PET) protocol, in which&#160;11C-AcAc and 18F-fluorodeoxyglucose (18F-FDG) are injected sequentially in each animal. This dual tracer PET acquisition is possible because of the short half-life of 11C (20.4 min). The rats also undergo a magnetic resonance imaging (MRI) acquisition seven days before the PET protocol. Prior to image analysis, PET and MRI images are coregistered to allow the measurement of regional cerebral uptake (cortex, hippocampus, striatum, and cerebellum). A quantitative measure of 11C-AcAc and 18F-FDG brain uptake (cerebral metabolic rate; &#956;mol/100 g/min) is determined by kinetic modeling using the image-derived input function (IDIF) method. Our new dual tracer PET protocol is robust and flexible; the two tracers used can be replaced by different radiotracers to evaluate other processes in the brain. Moreover, our protocol is applicable to the study of brain fuel supply in multiple conditions such as normal aging and neurodegenerative pathologies such as Alzheimer&#39;s and Parkinson&#39;s diseases.

Small-animal positron emission tomography enables the assessment of the brain&#39;s two main energy substrates:&#160;glucose and ketones. In the present method, 11C-acetoacetate and 18F-fluorodeoxyglucose are injected sequentially in each animal, and their uptake is measured quantitatively in specific brain regions determined from the magnetic resonance images.

We present a method for comparing the uptake of the brain&#39;s two key energy substrates: glucose and ketones (acetoacetate [AcAc] in this case) in the ...

An Engulfment Assay: A Protocol to Assess Interactions Between CNS Phagocytes and Neurons

Phagocytosis is a process in which a cell engulfs material (entire cell, parts of a cell, debris, etc.) in its surrounding extracellular environment and subsequently digests this material, commonly through lysosomal degradation. Microglia are the resident immune cells of the central nervous system (CNS) whose phagocytic function has been described in a broad range of conditions from neurodegenerative disease (e.g., beta-amyloid clearance in Alzheimer&#8217;s disease) to development of the healthy brain (e.g., synaptic pruning)1-6. The following protocol is an engulfment assay developed to visualize and quantify microglia-mediated engulfment of presynaptic inputs in the developing mouse retinogeniculate system7. While this assay was used to assess microglia function in this particular context, a similar approach may be used to assess other phagocytes throughout the brain (e.g., astrocytes) and the rest of the body (e.g., peripheral macrophages) as well as other contexts in which synaptic remodeling occurs (e.g. ,brain injury/disease).

Microglia are the resident immune cells of the central nervous system (CNS) with a high capacity to phagocytose or engulf material in their extracellular environment. Here, a broadly applicable, reliable, and highly quantitative assay for visualizing and measuring microglia-mediated engulfment of synaptic components is described.

Phagocytosis is a process in which a cell engulfs material (entire cell, parts of a cell, debris, etc.) in its surrounding extracellular environment and ...

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 ...

Preparation of Mouse Retinal Cryo-sections for Immunohistochemistry

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for determining the mechanisms underlying retinal diseases. Maintaining structural integrity throughout the tissue preparation is vital for obtaining reproducible retinal IHC data but can be challenging due to the fragility and complexity of retinal cytoarchitecture. Strong fixatives like 10% formalin or Bouin's solution optimally preserve the retinal structure, they often impede IHC analysis by enhancing the background fluorescence and/or diminishing antibody-epitope interactions, a process known as epitope masking. Milder fixatives, on the other hand, like 4% paraformaldehyde, reduces background fluorescence and epitope-masking, meticulous dissection techniques must be utilized to preserve the retinal structure. In this article, we present a comprehensive method to prepare mouse ocular posterior cups for IHC that is sufficient to preserve most antibody-epitope interactions without loss of retinal structural integrity. We include representative IHC with antibodies to various retinal cell type markers to illustrate tissue preservation and orientation under optimal and sub-optimal conditions. Our goal is to optimize IHC studies of the retina by providing a complete protocol from ocular posterior cup dissection to IHC.

This report describes comprehensive methods for preparing frozen mouse retina sections for immunohistochemistry (IHC). Methods described include dissection of the ocular posterior cup, paraformaldehyde fixation, embedding in Optimal Cutting Temperature (OCT) media and tissue orientation, sectioning and immunostaining.