This work is supported by the Australian Research council (ARC DP0988227) and the Biomedical Science Research Initiative Grant from the Discipline of Biomedical Science, The University of Sydney. The equipment Patch Clamp Amplifier EPC 8 was funded by the Startup Fund from the Discipline of Biomedical Science, The University of Sydney. The equipment InstruTECH LIH 8+8 Data Acquisition System was purchased with the funds from Rebecca L. Cooper Foundation and Startup Fund from the Discipline of Biomedical Science, The University of Sydney. We would like to thank the anonymous reviewers for their insightful suggestions and comments.

1. General Set Up and Tissue Preparation
<ol>
	<li>Keep the mouse in darkness for 30 min (aim to reduce its stress level). While waiting, prepare 1 L of extracellular solution. First dissolve 1.9 g of sodium bicarbonate in half a liter of Milli-Q water. Its pH is maintained at 7.4 by bubbling with 95% O2 and 5% CO2. Five minutes later, dissolve 8.8 g of Ames Medium in 100 ml of Milli-Q water, add to the sodium bicarbonate solution, top up to 1 L with Milli-Q water and mix well. Keep this solution carboxygenated for the rest of the experiment.</li>
	<li>Take 250 ml of the carboxygenated Ames Medium and set up the fluid reperfusion system in the electrophysiological set-up. Keep the solution carboxygenated.</li>
	<li>Uniformly smear a thin layer of vacuum grease onto the bottom of the perfusion chamber. Seal it with a cover slip and make sure it is air-tight. Take a small amount of grease (~ 2 mm in diameter) and position it on the top and bottom ends of the chamber. These balls will hold the grid (platinum frame strung with dental floss) in place and prevent it pressing too hard on the retina. Fix the chamber onto the platform and align both openings.</li>
	<li>The animal is first anaesthetized with Isoflurane for 2 min, and then sacrificed by cervical dislocation. Quickly remove eyes with a pair of small scissors (curved blades), wash thoroughly with carboxygenated extracellular fluid in a small beaker, and then place in a Petri dish filled with carboxygenated extracellular fluid.</li>
	<li>Under the dissecting microscope, make an insertion hole in the cornea (~ 2 mm from the edge) using a 19 gauge needle. Repeat for the other eye. This important step should be quick to allow oxygenation of both retinae. Remove one cornea with a small pair of iris scissors by cutting parallel to its edge. Next, remove the iris and lens with a pair of fine forceps. Repeat for the other eye. Transfer one eye cup into a beaker containing carboxygenated extracellular solution.</li>
	<li>With a scalpel blade, cut one eye cup into half, to be able to flatten the whole-mount and to obtain two whole-mounts per retina. Carefully separate the retina from the pigment epithelium using a blunt dissecting probe or by gently pulling it off. Once detached, the retina is fairly transparent. Note the convex surface is the photoreceptors side. The concave surface is the ganglion cells side and it may stick to itself due to the presence of sticky vitreous humour.</li>
	<li>With a pair of fine forceps, hold the edge of the detached retinal tissue close to the ora serrata and grab the ora serrata with another pair of fine forceps, pull it towards the center of the retina. This is a delicate process and may take a few attempts. If successful, ora serrata, ciliary body and vitreous humor are removed by this process, and the curvature of the retina is reduced (i.e. flatter).</li>
	<li>Trim the edges, and then transfer it to the perfusion chamber with ganglion cells facing up. Hold the tissue in place by placing the grid onto the grease balls. Place the remaining tissue with the other eye cup for later use. Similar procedures for retinal dissection are described in Middleton and Protti 16 and Pottek et al. 17.</li>
	<li>Mount the recording chamber under an upright microscope. Immediately set up the continuous perfusion of the retina with carboxygenated extracellular solution (35.5 &deg;C) at a rate of 3 - 5 ml/min. Make sure the grounding electrode is in place.</li>
	<li>Attached to the microscope is a digital camera allowing visualization of the tissue. The whole set up sits above an air table (shock absorption) and inside a Faraday cage (blocks external static and non-static electric fields).</li>
	<li>Turn on the light source (infrared, &gt; 850 nm) and with a 40X objective, bring its focus onto the cell bodies of ganglion cells. Find the central area of the whole-mount for recording.</li>
	<li>Defrost a frozen vial of K+ based intracellular solution (300 &mu;l, pH = 7.4) containing (in mM) K-gluconate: 140, HEPES acid: 10, MgCl2: 4.6, ATP-Na+: 4, GTP-Na+: 0.4 and EGTA: 10. All reagents purchased from Sigma-Aldrich.</li>
</ol>
2. Patching Cell Bodies of Retinal Ganglion Cells
<ol>
	<li>First fire polish the ends of borosilicate capillary glass tubing, then pull them with a Glass Microelectrode Puller with two heating steps (resistance: 5 - 8 M&Omega;).</li>
	<li>Add 3 &mu;l of Lucifer Yellow (final concentration 0.2%) to the intracellular solution, mix well, and centrifuge it. Next, transfer the top 85% of the solution into a 1 ml syringe, connect it to a 0.22 &mu;m filter unit, then to a 30 G reusable hypodermic needle. Refrigerate.</li>
	<li>Fix the glass pipette, of similar tip size as those used for recordings, in its holder and move it under the microscope using a micromanipulator. Under a 40X objective, slowly lower the pipette until it makes a small dimple on the surface of the inner limiting membrane. Carefully advance the pipette forward until it catches a small amount of tissue, and then move it upward and/or sideways to tear a small hole to expose the cell bodies of the ganglion cells. The smaller the hole, the higher the degree of integrity of the retinal network is maintained. If desired, sulforhodamine 101 (SR101; 0.5 - 1 &mu;M) can be added to the extracellular medium to label damaged neurons by using fluorescence microscopy 15.</li>
	<li>Fill up a clean pipette with 8 - 10 &mu;l of intracellular solution. Insert it into the holder attached to the amplifier's head stage and make sure the chlorinated silver chloride electrode is submerged. Discard if dirt and/or air bubbles is/are present.</li>
	<li>Turn on all equipment. We used an EPC 8 patch clamp amplifier controlled by PatchMaster to achieve whole-cell recordings in voltage clamp configuration. Note that other amplifiers that allow current clamp recordings in fast mode can also be used. Apply and maintain positive pressure in the pipette solution. Select a cell with a large cell body so most likely it is a ganglion cell rather than a displaced amacrine cell or a glial cell. Slowly lower the pipette tip so it is making a small dimple on the surface of the membrane, stop, and release the pressure. Immediately lower the holding potential to - 60 mV. Once a gigaseal (G&Omega;) is achieved between them, gently suck to break the membrane to achieve a whole-cell patch clamp configuration. This procedure is a delicate step, if too much negative pressure is applied, then the connection becomes leaky; if too little, the membrane remains intact.</li>
	<li>Over time, Lucifer Yellow will fill the cell, allowing visualization of the cell's morphology. If stable, this set-up should last 30 min or more.</li>
</ol>
3. Recordings of Ganglion Cells Using Dynamic Clamp
<ol>
	<li>First, a current-voltage test (from - 75 to + 35 mV every 10 mV) is executed using PatchMaster. Proceed to later tests only if a large Na+ current (&gt; 1 nA) is present. Note that in the rare occasion where a displaced amacrine cell is patched, in that case, will not respond with trains of action potentials to subsequent current injections.</li>
	<li>Open LabVIEW and execute the custom written Neuroakuma program, then load various conductance waveforms. Set repeat number to 8 (6 - 8 for statistical purpose). Set reversal potentials of excitatory and inhibitory conductances at 0 and - 75 mV respectively. Select one matching pair of excitatory and inhibitory conductances for testing. Excitatory conductance trace (in green) and inhibitory conductance trace (in red) will appear in the lower panel.</li>
	<li>Go back to PatchMaster, select current clamp mode, and immediately switch the amplifier to 'CC + Comm FAST' current clamp mode. This mode allows to accurately follow rapid changes in membrane potential. If a good seal between the membrane and pipette is maintained, little/no compensation is needed; otherwise, the recording will not last long. In Neuroakuma, press the "record" button. The cellular response (usually with a phasic burst of action potentials, Figure 1A) will appear on the upper panel. Inject current into the cell with low scaling of conductances first, if little/no response is observed, increase in small increments until it produces a strong response. Excessive current injection can kill the cell. Once tests are completed, select a new pair of conductance waveforms, repeat the above for all pairs of physiological and artificial conductances. The physiological currents (I) injected in real time are based on: 
	I(t) = Gexc(t) x (Vm(t) - Vexc) + Ginh(t) x (Vm(t) - Vinh)</li>
</ol>
Conductance (G in nS) could be synaptic or artificial. Excitatory and inhibitory synaptic conductance waveforms were collected from previous experiments performed by Protti, Di Marco, Huang, Vonhoff, Nguyen and Solomon (unpublished results) in response to different visual stimuli in control conditions and in the presence of tetrodotoxin (TTX, 1 &mu;M). Artificial conductance waveforms were modeled using an alpha function. Vm (in mV) is the recorded membrane potential. Gexc and Ginh represent excitatory and inhibitory conductances respectively whilst Vexc and Vinh represent the reversal potentials of excitatory and inhibitory conductances respectively. Time is t in ms. Sampling rate is 40 kHz. 
Is = I0(t/&alpha;)e-&alpha;t
The above equation is an alpha function (I0 = maximum current; 1/&alpha; = time to peak (sec-1) of the current). The rise time and decay time of the synaptic current is dictated by &alpha; 4. The excitatory conductance was unchanged whilst the latency of the inhibitory conductance was modified, reducing its delay relative to the onset of excitation (Figure 2D).
<ol start="4">
	<li>After recording, carefully retract the pipette so the cell body stays intact. Immediately fix the tissue in 4% paraformaldehyde for 30 min, followed by three 10 min washes with 0.1 M of phosphate buffer. Refrigerate and perform antibody staining the following day or within a week.</li>
</ol>
4. Antibody Staining Against Lucifer Yellow Filled Retinal Ganglion Cells
<ol>
	<li>Antibody staining against Lucifer Yellow filled ganglion cells at room temperature (primary anti-Lucifer Yellow rabbit IgG (dilution = 1:10,000) for five days; on the sixth day, overnight staining with secondary goat anti-rabbit IgG (dilution = 1:500)). Wet mount the retina in Fluorescent Preserving Media. Cover slip and seal with nail polish.</li>
	<li>Take confocal pictures of the cell morphology using a Leica Spec-II confocal microscope (Figure 1B). The presence of an axon allows the confirmation of ganglion cells.</li>
</ol>

<ol>
	<li>Robinson, H. P. C., Kawai, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Injection+of+digitally+synthesized+synaptic+conductance+transients+to+measure+the+integrative+properties+of+neurons.">Injection of digitally synthesized synaptic conductance transients to measure the integrative properties of neurons.</a> Journal of Neuroscience Methods. 49, 157-165 (1993).</li><li>Sharp, A. A., O'Neil, M. B., Abbott, L. F., Marder, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+dynamic+clamp:+artificial+conductances+in+biological+neurons.">The dynamic clamp: artificial conductances in biological neurons.</a> Trends in Neuroscience. 16, 389-394 (1993).</li><li>Sharp, A. A., O'Neil, M. B., Abbott, L. F., Marder, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+clamp:+computer-generated+conductances+in+real+neurons.">Dynamic clamp: computer-generated conductances in real neurons.</a> Journal of Neurophysiology. 69, 992-995 (1993).</li><li>Johnson, D., Wu, S. M. -. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Foundations of cellular neurophysiology. , (1995).</li><li>Taylor, W. R., 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=Diverse+synaptic+mechanisms+generate+direction+selectivity+in+the+rabbit+retina.">Diverse synaptic mechanisms generate direction selectivity in the rabbit retina.</a> Journal of Neuroscience. 22 (17), 7712-7720 (2002).</li><li>Jurkat-Rott, K., Lehmann-Horn, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+patch+clamp+technique+in+ion+channel+research.">The patch clamp technique in ion channel research.</a> Current Pharmaceutical Biotechnology. 4, 221-238 (2003).</li><li>Prinz, A. A., Abbott, L. F., Marder, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+dynamic+clamp+comes+of+age.">The dynamic clamp comes of age.</a> Trends in Neuroscience. 27, 218-224 (2004).</li><li>Williams, S. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+compartmentalization+and+functional+impact+of+conductance+in+pyramidal+neurons.">Spatial compartmentalization and functional impact of conductance in pyramidal neurons.</a> Nature Neuroscience. 7 (9), 961-967 (2004).</li><li>Perkins, K. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-attached+voltage-clamp+and+current-clamp+recording+and+stimulation+techniques+in+brain+slices.">Cell-attached voltage-clamp and current-clamp recording and stimulation techniques in brain slices.</a> Journal of Neuroscience Methods. 154 (1-2), 1-18 (2006).</li><li>Feng, S. S., Jaeger, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+SK+calcium-dependent+potassium+currents+in+regulating+the+activity+of+deep+cerebellar+nucleus+neurons:+a+dynamic+clamp+study.">The role of SK calcium-dependent potassium currents in regulating the activity of deep cerebellar nucleus neurons: a dynamic clamp study.</a> Cerebellum. 7 (4), 542-546 (2008).</li><li>Butera, R., Lin, R., Destexhe, A., Bal, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Key+factors+for+improving+dynamic+clamp+performance.+In:+Dynamic-Clamp+From+Principles+to+Applications+Series.">Key factors for improving dynamic clamp performance. In: Dynamic-Clamp From Principles to Applications Series.</a> Dynamic-Clamp From Principles to Applications Series: Springer Series in Computational Neuroscience. 1, (2009).</li><li>Marco, S. D. i., Nguyen, V. A., Bisti, S., Protti, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Permanent+functional+reorganization+of+retinal+circuits+induced+by+early+long-term+visual+deprivation.">Permanent functional reorganization of retinal circuits induced by early long-term visual deprivation.</a> The Journal of Neuroscience. 29 (43), 13691-13701 (2009).</li><li>Zhao, Y., Inayat, S., Dikin, D. A., Singer, J. H., Ruoff, R. S., Troy, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patch+clamp+technique:+review+of+the+current+state+of+the+art+and+potential+contributions+from+nanoengineering.">Patch clamp technique: review of the current state of the art and potential contributions from nanoengineering.</a> Proceedings of the Institution of Mechanical Engineers, Part N: Journal of Nanoengineering and Nanosystems. 222 (1), 1-11 (2009).</li><li>Lobb, C. J., Paladini, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+a+NMDA+receptor+conductance+in+rat+midbrain+dopaminergic+neurons+using+the+dynamic+clamp+technique.">Application of a NMDA receptor conductance in rat midbrain dopaminergic neurons using the dynamic clamp technique.</a> J. Vis. Exp. (46), e2275 (2010).</li><li>Briggman, K. L., Euler, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bulk+electroporation+and+population+calcium+imaging+in+the+adult+mammalian+retina.">Bulk electroporation and population calcium imaging in the adult mammalian retina.</a> Journal of Neurophysiology. 105 (5), 2601-2609 (2011).</li><li>Middleton, T. P., Protti, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cannabinoids+modulate+spontaneous+synaptic+activity+in+retinal+ganglion+cells.">Cannabinoids modulate spontaneous synaptic activity in retinal ganglion cells.</a> Visual Neuroscience. 28 (5), 393-402 (2011).</li><li>Pottek, M., Knop, G. C., Weiler, R., Dedek, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+characterization+of+GFP-expressing+cell+populations+in+the+intact+retina.">Electrophysiological characterization of GFP-expressing cell populations in the intact retina.</a> J. Vis. Exp. (57), e3457 (2011).</li></ol>

All procedures were first approved by the Animal Care Ethics Committee of The University of Sydney, and then performed in accordance with the guidelines of the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes (National Health and Medical Research Council of Australia).

Here we show the use of dynamic clamp to assess the influence of the ratio and relative timing of excitation and inhibition on retinal ganglion cell output. Dynamic clamp makes use of computer simulations to introduce physiologically recorded or artificial synaptic conductances into living neurons. This methodology provides an interactive tool by which conductances can be modified and injected into neurons for computing their influence on neuronal responses. Conductance waveforms can be obtained from experiments in which...

The retina is a near-transparent neural tissue lining the back of the eye. Many studies use the retina as the model to investigate the first steps in visual processing and mechanisms of synaptic signaling. Since the retinal network in the whole-mount preparation remains intact after dissection, it represents an ideal system to study synaptic interactions as its physiological responses are very similar to the in vivo conditions. Thus, using an isolated retina the properties of its neurons can be studied using patch clamp techniques (for reviews on the technique, see 6,9,13). Identification of the exact contribution of specific circuits and neurotransmitters to ganglion cell response, however, is usually hindered as pharmacological agents act on various sites.
Physiological responses of retinal neurons to light, the natural stimulus, can be recorded with glass pipettes filled with intracellular fluid. Using patch clamp techniques, neuronal responses to light stimulation can be recorded as membrane potential fluctuations (current clamp) or as currents (voltage clamp). By holding the membrane potential at different voltages and implementing a posteriori conductance analysis, it is possible to isolate inhibitory and excitatory synaptic inputs 5,12. This type of experiments can be carried out in normal bathing medium and in the presence of different pharmacological agents to isolate the contribution of different neurotransmitters and receptors to neuronal responses. A wealth of studies from many laboratories characterized the dependence of spiking output and excitatory and inhibitory inputs on stimulus properties such as size, contrast, spatial and temporal frequencies, direction, orientation and other stimulus variables. Although these experimental approaches provide information about the relationship between spike output and synaptic inputs as a function of stimulus properties, interpretation of the contribution of specific cell types and their synaptic inputs to cell excitability is not straightforward. This is due to the fact that typically both excitatory and inhibitory inputs vary with stimulus properties and thus, it is not possible to assess the precise impact that changes in either of these inputs has on neuronal spiking.
An alternative approach to circumvent these limitations is to carry out dynamic clamp recordings, which allow a critical evaluation of the contribution of individual synaptic inputs to spiking output. The dynamic clamp technique allows direct injection of current into the cell and the amount of current injected at a given time depends on the recorded membrane potential at that time 1,2,3 (for review, see 7,14). It is a modified current clamp set-up where a real-time, fast feedback interaction between the cell under recording and the equipment comprising specialized hardware, software and a computer is achieved. The amount of current injected into the cell is computed accordingly. Hence, the advantage of this method is that the cell can be stimulated with different combinations of conductance waveforms, and its response will mimic the activation of receptors that mediate synaptic inputs. For example, comparison of the response to injection of excitatory and inhibitory conductances for a small spot with the response to injection of excitatory conductance for a small spot only provides information about the impact of inhibition on cell response. Likewise, other combinations of physiologically recorded conductances can be co-injected to reveal how stimulus-dependent changes in excitatory and/or inhibitory conductances affect spike output.
In our study, the dynamic clamp technique is used to demonstrate the impact of the relative amplitude and timing of synaptic inputs on the firing properties of retinal ganglion cells. Various conductances obtained from physiological experiments in control conditions or in the presence of pharmacological agents were employed as inputs. In addition, artificial conductances based on alpha functions were also used in order to investigate how synaptic inputs are integrated by neurons. Thus this is a versatile technique that allows various types of conductance generated either physiologically, pharmacologically or computationally to be injected into the same ganglion cell, so comparison of responses to these inputs can be made.

<table border="1">
	<tbody>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Isoflurane Inhalation Anaesthetic</td>
			<td>Pharmachem</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Ames Medium with L-Glutamate (Powder)</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Potassium Gluconate, Anhydrous</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>HEPES Sodium salt</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Magnesium chloride solution (4.9 mol/l)</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Adenosine 5'-triphosphate (ATP) disodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Guanosine 5'-triphosphate sodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Ethylene glycol-bis(2-aminoethylether)-N,N,N',N'-tetraacetic acid</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Paraformaldehyde Powder, 95%</td>
			<td>Sigma-Aldrich</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Anti-Lucifer Yellow, Rabbit IgG Fraction (3 mg/ml)</td>
			<td>Invitrogen</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Goat Anti-Rabbit IgG (H+L) 2 mg/ml</td>
			<td>Invitrogen</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Fluorescent Preserving Media</td>
			<td>BioFX Laboratories Inc.</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Capillary Glass Tubing with flame polished ends (OD = 1.50 mm, ID = 0.86 mm, Length = 15 cm)</td>
			<td>Warner Instruments</td>
			<td>64-0794</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Single Stage Glass Microelectrode Puller</td>
			<td>Narishinge Japan</td>
			<td>Model PP-830</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Minipuls 2</td>
			<td>Gilson</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Millex-GV 0.22 &mu;m Filter Unit</td>
			<td>Millipore Corporation</td>
			<td>SLGV004SL</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Luer Lock Reusable Hypodermic Needle: 30 G</td>
			<td>Smith &amp; Nephew (Australia)</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Single Inline Solution Heater</td>
			<td>Warner Instruments</td>
			<td>Model SH-27B</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dual Automatic Temperature Controller</td>
			<td>Warner Instruments</td>
			<td>TC-344B</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Olympus Stereomicroscope SZ61</td>
			<td>Olympus Corporation</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Olympus Microscope BX50WI: with 40X objective</td>
			<td>Olympus Corporation</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>0-30 V 2.5 A DC Power Supply</td>
			<td>Dick Smith Electronics</td>
			<td>Q1770</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Digital Microscopic Camera ProgResMF cool</td>
			<td>Jenoptik</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Micromanipulator MP-225</td>
			<td>Sutter Instrument Company</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Patch Clamp Amplifier EPC 8</td>
			<td>HEKA Elektronik</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>InstruTECH LIH 8+8 Data Acquisition System</td>
			<td>HEKA Elektronik</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Computer: DELL</td>
			<td>Dell Corporation</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
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implementing dynamic clamp with synaptic and artificial conductances in mouse retinal ganglion cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

The contribution of different sources of inhibitory inputs to ganglion cell responses is demonstrated through the application of various conductance waveforms. These waveforms were obtained with stimuli of different luminance in normal conditions and in the presence of TTX, a voltage-gated Na+ channel blocker that blocks action potential generation only in a subset of inhibitory retinal interneurons. Figure 2A shows a representative response to injection of excitatory and inhibitory conductanc...

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Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

implementing-dynamic-clamp-with-synaptic-artificial-conductances

Research

JoVE Journal

Neuroscience

12.2K 次观看.  University of Sydney. 此视频文章说明设置，程序，修补细胞组织，以及如何实现动态钳记录整个安装鼠标视网膜神经节细胞的。这种技术允许调查的兴奋性和抑制性突触输入精确的贡献，他们的相对幅度和时机神经扣球。

视频: Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

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 ...

An Isolated Retinal Preparation to Record Light Response from Genetically Labeled Retinal Ganglion Cells

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1. This information is then segregated into what are known as the ON and OFF pathways. The photoreceptors signal light information to the bipolar cells (BCs), which depolarize in response to increases (On BCs) or decreases (Off BCs) in light intensity. This segregation of light information is maintained at the level of the retinal ganglion cells (RGCs), which have dendrites stratifying in either the Off sublamina of the inner plexiform layer (IPL), where they receive direct excitatory input from Off BCs, or stratifying in the On sublamina of the IPL, where they receive direct excitatory input from On BCs. This segregation of information regarding increases or decreases in illumination (the On and Off pathways) is conserved and signaled to the brain in parallel.
The RGCs are the output cells of the retina, and are thus an important cell to study in order to understand how light information is signaled to visual nuclei in the brain. Advances in mouse genetics over recent decades have resulted in a variety of fluorescent reporter mouse lines where specific RGC populations are labeled with a fluorescent protein to allow for identification of RGC subtypes 2 3 4 and specific targeting for electrophysiological recording. Here, we present a method for recording light responses from fluorescently labeled ganglion cells in an intact, isolated retinal preparation. This isolated retinal preparation allows for recordings from RGCs where the dendritic arbor is intact and the inputs across the entire RGC dendritic arbor are preserved. This method is applicable across a variety of ganglion cell subtypes and is amenable to a wide variety of single-cell physiological techniques.

This article provides a description of how to dissect and record from the isolated retinal preparation in mouse. In particular, we describe how to record light responses from a fluorescently labeled ganglion cell population and subsequently identify and analyze its morphology.

The first steps in vertebrate vision take place when light stimulates the rod and cone photoreceptors of the retina 1.  This information is then ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. Retinal neurons in the network comprise of numerous subtypes. More than 10 subtypes of bipolar cells, ganglion cells, and amacrine cells have been identified by morphological studies. Multiple subtypes of retinal neurons are thought to encode distinct features of visual signaling, such as motion and color, and form multiple neural pathways. However, the functional roles of each neuron in visual signal processing are not fully understood. The patch clamp method is useful to address this fundamental question. Here, a protocol to record light-evoked synaptic responses in mouse retinal neurons using patch clamp recordings in dark-adapted conditions is provided. The mouse eyes are dark-adapted O/N, and retinal slice preparations are dissected in a dark room using infrared illumination and viewers. Infrared light does not activate mouse photoreceptors and thus preserves their light responsiveness. Patch clamp is used to record light-evoked responses in retinal neurons. A fluorescent dye is injected during recordings to characterize neuronal morphological subtypes. This procedure enables us to determine the physiological functions of each neuron in the mouse retina.

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. ...

Single-cell RNA-Seq of Defined Subsets of Retinal Ganglion Cells

The discovery of cell type-specific markers can provide insight into cellular function and the origins of cellular heterogeneity. With a recent push for the improved understanding of neuronal diversity, it is important to identify genes whose expression defines various subpopulations of cells. The retina serves as an excellent model for the study of central nervous system diversity, as it is composed of multiple major cell types. The study of each major class of cells has yielded genetic markers that facilitate the identification of these populations. However, multiple subtypes of cells exist within each of these major retinal cell classes, and few of these subtypes have known genetic markers, although many have been characterized by morphology or function. A knowledge of genetic markers for individual retinal subtypes would allow for the study and mapping of brain targets related to specific visual functions and may also lend insight into the gene networks that maintain cellular diversity. Current avenues used to identify the genetic markers of subtypes possess drawbacks, such as the classification of cell types following sequencing. This presents a challenge for data analysis and requires rigorous validation methods to ensure that clusters contain cells of the same function. We propose a technique for identifying the morphology and functionality of a cell prior to isolation and sequencing, which will allow for the easier identification of subtype-specific markers. This technique may be extended to non-neuronal cell types, as well as to rare populations of cells with minor variations. This protocol yields excellent-quality data, as many of the libraries have provided read depths greater than 20 million reads for single cells. This methodology overcomes many of the hurdles presented by Single-cell RNA-Seq and may be suitable for researchers aiming to profile cell types in a straightforward and highly efficient manner.

Here, we present a combinatorial approach for classifying neuronal cell types prior to isolation and for the subsequent characterization of single-cell transcriptomes. This protocol optimizes the preparation of samples for successful RNA Sequencing (RNA-Seq) and describes a methodology designed specifically for the enhanced understanding of cellular diversity.

The discovery of cell type-specific markers can provide insight into cellular function and the origins of cellular heterogeneity. With a recent push for ...

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in vivo&#8212;rapidly, repetitively, and at a high resolution. This protocol describes OCT imaging in the mouse retina as a powerful tool to study optic neuropathies (OPN). The OCT system is an interferometry-based, non-invasive alternative to common post mortem histological assays. It provides a fast and accurate assessment of retinal thickness, allowing the possibility to track changes, such as retinal thinning or thickening. We present the imaging process and analysis with the example of the Opa1delTTAG mouse line. Three types of scans are proposed, with two quantification methods: standard and homemade calipers. The latter is best for use on the peripapillary retina during radial scans; being more precise, is preferable for analyzing thinner structures. All approaches described here are designed for retinal ganglion cells (RGC) but are easily adaptable to other cell populations. In conclusion, OCT is efficient in mouse model phenotyping and has the potential to be used for the reliable evaluation of therapeutic interventions.

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

This manuscript describes a protocol for the in vivo imaging of the mouse retina with high-resolution spectral domain optical coherence tomography (SD-OCT). It focuses on retinal ganglion cells (RGC) in the peripapillary region, with several scanning and quantifying approaches described.

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in ...

Partial Optic Nerve Transection in Rats: A Model Established with a New Operative Approach to Assess Secondary Degeneration of Retinal Ganglion Cells

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection is considered a useful and reproducible model. Compared with other optic nerve injury models used commonly for assessing secondary degeneration, e.g. complete optic nerve transection and optic nerve crush models, the partial optic nerve transection model is superior as it distinguishes primary from secondary degeneration in situ. Therefore, it serves as an excellent tool for evaluating secondary degeneration. This study describes a novel operative approach of partial optic nerve transection by directly accessing the area of the retrobulbar optic nerve through the orbital lateral wall of the eyeball. Moreover, we present a newly designed, low cost surgical instrument to assist with transection. As demonstrated by the representative results in distinguishing the boundary of primary and secondary injury areas, the new approach and instrument ensures high efficiency and stability of the model by providing adequate space for surgical operation. This in turn makes it easy to separate the meningeal sheath and ophthalmic vessels from the optic nerve before transection. An additional benefit is that this space-saving operative approach improves the investigators&#39; ability to administer drugs, carriers, or selective RGC tracers to the stump of the partially transected optic nerve, allowing the exploration of mechanisms behind secondary injury in RGCs, in a new way.

Secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. This study describes an innovative operative approach for partial optic nerve transection. The use of this space-saving operative approach extends the model's application range, and allows exploration of secondary injury mechanisms in RGCs in a new way.

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...

Evaluation of Synaptic Multiplicity Using Whole-cell Patch-clamp Electrophysiology

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic multiplicity). Synaptic multiplicity is plastic and changes throughout development and in different physiological conditions, being an important determinant for the efficacy of synaptic transmission. Here, we outline experiments for estimating the degree of multiplicity of synapses terminating onto a given postsynaptic neuron using whole-cell patch clamp electrophysiology in acute brain slices. Specifically, voltage-clamp recording is used to compare the difference between the amplitude of spontaneous excitatory postsynaptic currents (sEPSCs) and miniature excitatory postsynaptic currents (mEPSCs). The theory behind this method is that afferent inputs that exhibit multiplicity will show large, action potential-dependent sEPSCs due to the synchronous release that occurs at each synaptic contact. In contrast, action potential-independent release (which is asynchronous) will generate smaller amplitude mEPSCs. This article outlines a set of experiments and analyses to characterize the existence of synaptic multiplicity and discusses the requirements and limitations of the technique. This technique can be applied to investigate how different behavioral, pharmacological or environmental interventions in vivo affect the organization of synaptic contacts in different brain areas.

Here, we present a protocol for evaluating the functional synaptic multiplicity using whole-cell patch clamp electrophysiology in acute brain slices.

In the central nervous system, a pair of neurons often form&#160;multiple synaptic contacts and/or functional neurotransmitter release sites (synaptic ...