1. Cell Dissociation
<ol>
	<li>A flowchart outlining the protocol is shown is Figure 1. For the catalog numbers of the particular reagents used throughout this protocol, please refer to Table 1. Dissect the retina in a PBS bath. During the dissection, it is best to remove the vitreous and the lens since keeping them with the retina can impede the dissociation. It is not always critically important to remove all of the retinal pigment epithelium (RPE) and in some instances it may be impossible to completely remove it. However, for single cell profiling experiments of photoreceptors, the RPE should be removed. Failure to remove the RPE can lead to contamination of the rod cell profiles with RPE expressed genes. This is presumably due to the connection between the photoreceptors and the RPE. 
	For the dissociation, whole adult murine retinas are incubated at 37 &deg;C for 10 min in a 1.5 ml microcentrifuge tube containing 20 &mu;l activated papain, 20 &mu;l 25 mM Cysteine in 5 mM EDTA, and 360 &mu;l Hank's Balanced Salt Solution (HBSS) supplemented with 10 mM HEPES. Filter-tipped pipette tips are used for all steps throughout the protocol to minimize potential contamination. The papain concentration and the time of incubation will vary according to the age and the nature of the tissue. For example, to dissociate developing retinas isolated at postnatal day 0 (P0), incubate the retina in 10 &mu;l activated papain, 10 &mu;l 25 mM Cysteine in 5 mM EDTA, and 380 &mu;l Hank's Balanced Salt Solution (HBSS) supplemented with 10 mM HEPES for 10 min at 37 &deg;C. Retinas from different species also require slightly different conditions. Chicken retinas are thinner than mouse retinas at most stages of development. Using 10 &mu;l activated papain, 10 &mu;l 25 mM Cysteine in 5 mM EDTA, and 380 &mu;l Hank's Balanced Salt Solution (HBSS) supplemented with 10 mM HEPES for 5 min at 37 &deg;C is sufficient to dissociate these retinas.</li>
	<li>Gently tap the tube to dislodge any settled cells, then triturate gently 10-20 times with a p1000 pipettor. Incubate for an additional 10 min at 37 &deg;C if large clumps of tissue persist and then re-triturate 10-20 times.</li>
	<li>Add 5 &mu;l of DNase I (10 U/&mu;l) and incubate for 5 min at room temperature. Triturate gently with a p1000 pipettor. The exact number of times will need to be determined empirically, but generally falls between 10-20.</li>
	<li>Centrifuge in a table-top centrifuge at 3000 rpm for 3 min Remove the supernatant, leaving 100-200 &mu;l of liquid at the bottom, and tap the tube to dislodge the pellet. Add 1 ml of HBSS and resuspend the pellet with gentle trituration. Tapping the tube is critical here as simply resuspending the pellet with lyse many of the retinal cells. This is especially true with adult murine retinas.</li>
	<li>Centrifuge at 3000 rpm for 3 min Carefully remove the supernatant and resuspend in 450 &mu;l of PBS containing 0.1% BSA. It is important to remove as much of the supernatant as possible to minimize the presence of mRNAs from lysed cells and any products from the dissociation that could inhibit future reactions.</li>
</ol>
2. Harvesting Single Cells
<ol>
	<li>Prepare two 6 cm dishes containing 5 ml of PBS supplemented with 0.1% BSA. Plate the dissociated cells onto one dish and allow the cells to settle for at least 5 min. The cell density used depends greatly on the nature of the single cells being profiled. For example, for very rare cells labeled with a fluorescent marker such a GFP, entire dissociated retinas are plated on a single 6 cm dish. For cells that are somewhat more abundant, fewer cells are initially plated to make it easier to harvest one (or very close to one) cell with the first micropipette.</li>
	<li>Place the plate of dissociated cells on an inverted microscope. We use the IMT-2 model from Olympus. Using micropipettes that have been drawn to a fine tip (inner diameter 0.5 mm, outer diameter 1.04 mm) (Figure 2) and together with an aspirator tube, harvest a single cell. The cells readily enter the micropipette when it is placed close to them on the dish. It is critical that the aspirator tube is neither too long nor too short for the distance to the stage associated with the inverted microscope (The aspirator tubes we use with our inverted microscope are 38.1 cm long). If the tube is too long, cells may not be expelled efficiently from the micropipette into the collection tube. The primary reason we use the older IMT-2 model inverted microscope is that we have found it has the ideal distance to the stage for our aspirator assembly tubes. After entering the micropipette, the cell (or cells) is expelled onto a second plate (wash plate) to ensure that one and only one cell is harvested for gene expression profiling. On the second plate it is often necessary to either remove extraneous cells with a new micropipette or move the cell of interest to a different location to ensure that only a single cell enters the micropipette.</li>
	<li>When an individual cell is completely isolated from neighboring cells, use a new micropipette to aspirate the cell of interest as in section 2.2 and then expel it directly into a 0.2 ml PCR tube containing 4.5 &mu;l cell lysis buffer. The cell is expelled onto the side of the tube, being careful not to break off the tip of the micropipette into the tube. The samples can be spun briefly in a bench top microcentrifuge to immerse the cell in lysis buffer. This spinning is usually done after every 5th cell and then again at the end of collection.Tubes containing single cells in lysis buffer should be kept on ice for the duration of the single cell isolation. For optimal results, single cells are collected within a two-hour window post-dissociation. After this time has elapsed, it is best to proceed to the reverse transcription step since additional time has been observed to increase the chances of RNA degradation.</li>
</ol>
3. Reverse Transcription
<ol>
	<li>Briefly spin the samples in a benchtop minicentrifuge to ensure all the single cells are immersed in the cell lysis buffer. To promote cell lysis, incubate the sample at 70 &deg;C for 90 sec. in a thermocycler. Immediately place the tubes back on ice.</li>
	<li>Leave the tubes on ice for 2 min. For all reagent additions, use filter-tip pipette tips to prevent contamination. To perform the reverse transcription, add 0.33 &mu;l Superscript III (200 U/&mu;l), 0.05 &mu;l RNase Inhibitor (40 U/&mu;l), and 0.07 &mu;l T4 gene 32 protein. Incubate the reaction mixture for 50 min at 50 &deg;C in a thermocycler. Inactivate the enzyme at 70 &deg;C for 15 min and place on ice.</li>
	<li>To remove the free primer, add 0.1 &mu;l Exonuclease Buffer (10X), 0.8 &mu;l dH20 (molecular biology grade), and 0.1 &mu;l Exonuclease I (20 U/&mu;l). Incubate at 37 &deg;C for 30 min in a thermocycler. Inactivate the enzyme by incubating the reaction at 80 &deg;C for 25 min and then placing the tubes on ice.</li>
</ol>
4. Tailing and Single-Cell PCR
<ol>
	<li>Add 6 &mu;l of the tailing reaction mixture and use a thermocycler to incubate the sample at 37 &deg;C for 20 min, 70 &deg;C for 10 min, and hold at 4 &deg;C</li>
	<li>Add 10 &mu;l of the tailed sample to the PCR reaction mixture and perform the second strand synthesis and PCR amplification using the following conditions:
	<ol>
		<li>95 &deg;C for 2 min</li>
		<li>37 &deg;C for 5 min</li>
		<li>72 &deg;C for 16 min</li>
		<li>93 &deg;C for 40 sec</li>
		<li>67 &deg;C for 1 min</li>
		<li>72 &deg;C for 6 min, adding 6 sec per cycle</li>
		<li>Go to step 4 34 times</li>
		<li>72 &deg;C for 10 min</li>
		<li>Hold at 4 &deg;C</li>
	</ol>
	</li>
</ol>
5. Labeling for Affymetrix Chips
<ol>
	<li>Label 10-20 &mu;g of cDNA (the concentration of the amplified cDNA resulting from a single cell is usually ~1 &mu;g/&mu;l) to obtain a robust hybridization on Affymetrix microarrays. First, fragment the cDNA by adding 10-20 &mu;l of the single-cell cDNA to 8 &mu;l 1X One-Phor-All buffer and 1 &mu;l of diluted (1:10 in 1X One-Phor-All buffer) DNaseI in an 80 &mu;l reaction. Incubate in a thermocycler for 13 min at 37 &deg;C. Inactivate the DNase I for 15 min at 100 &deg;C and place on ice.</li>
	<li>Add 20 &mu;l 5X TdT buffer, 2.5 &mu;l Biotin N6-ddATP (Enzo Biosciences), and 1 &mu;l TdT (diluted 1:8 in TdT buffer).</li>
	<li>Incubate for 90 min at 37 &deg;C, then 5 min at 65 &deg;C. Store at -20 &deg;C or hybridize immediately to an Affymetrix microarray. The hybridization is performed using standard Affymetrix protocols.</li>
</ol>
6. Representative Results
To assess the quality of the cDNA, 10 &mu;l of the cDNA sample was loaded on a 1% agarose gel. The cDNA is mainly judged by the size range (500-2000 bp) and intensity of the cDNA smear (Figure 1 and Figure 3a). In the gel depicted in Figure 1, every other lane shows a cDNA smear from retinal cells isolated E16.5. The lanes in between show the resulting smears when only media is aspirated into the needle and deposited into the PCR tube. These lanes are never completely devoid of a faint smear, but they do not show any results when gene-specific PCR primers are used to amplify genes from them. Additionally, the intensity of the cDNA smear can vary somewhat (compare Figure 1 and Figure 3a). In Figure 3a lanes a, f, g, and h contain robust cDNA smears while lanes b, c, d, and e are considerably less robust.
Gene-specific PCR is used as a secondary screen of the quality of the cDNA from a single cell. The cDNAs in Figure 3 were isolated from fluorescent retinal ganglion cells and they were tested using PCR primers for the retinal ganglion cell markers Brn3b and Pax6. For this assay, 1 &mu;l of the cDNA was subjected to PCR for 30 cycles. Real-time quantitative PCR would be a preferable assay, but it can be prohibitively expensive and is not necessary for merely assessing the cDNA quality. All 8 of the single cells tested positive for Brn3b and 6 out 8 for Pax6 (Figure 3b). Even the cDNA smears that were less robust produced bands for Brn3b and Pax6. Despite these PCR results, it is our experience that cDNA smears such as those in lanes b, c, d, and e do not yield robust hybridizations on Affymetrix arrays and are generally avoided. Gene specific PCR for the photoreceptor marker Crx was used to determine the level of contamination in the single cell cDNAs (Figure 3b). A photoreceptor marker was chosen because these cells make up the majority of the retina (~70%) and, therefore, any cell lysis that occurred would most likely involve rod photoreceptors. There was only a faint amount of Crx present in these cDNA preparations even after 30 cycles of PCR. Finally, this PCR-based screen can be used to begin to determine which subsets of a particular cell type the cDNAs were derived from before subjecting them to a full microarray profiling. This can help to prioritize the cells that are profiled, as this is the most expensive step in the process. For example, we have identified subsets of retinal ganglion cells by screening them for the presence of the gene Tachykinin1 (Figure 3b).
From the small PCR-based screen, particular single cells are chosen and their cDNAs hybridized to an Affymetrix microarray. The microarray data is compared and patterns can be discovered using standard clustering algorithms. Heatmaps, such as in Figure 4, are generated to graphically represent the data. There are two main conclusions regarding the single cell profiling data in Figure 4. First, genes expressed in specific cell types are found almost exclusively in those cell types after the single cell protocol is performed. In most instances where marker genes of one cell type are found also in a second cell type, such as the observation that bipolar cells express some "rod" genes at lower levels, this expression in the second cell type is readily confirmed by either in situ hybridization or antibody staining. Second, within the more diverse amacrine and ganglion cell profiles, heterogeneity of gene expression is immediately apparent. Pou4f1 is only expressed in about 1/4 of the developing ganglion cells, while Nr4a2 and Scube2 are two examples of heterogeneously expressed genes in different amacrine cells. In fact, the genes shown in the heatmap are just a small sample as several hundred genes have been identified and confirmed as markers of developing ganglion cells or as markers of different populations of amacrine cells2,4.
<img alt="Figure 1" src="/files/ftp_upload/3824/3824fig1.jpg" /> 
Figure 1. Flowchart of the single cell profiling method. First retinas are isolated and dissociated into individual cells using papain. Second, single cells are harvested with a pulled glass needle and deposited into PCR tubes. The cells are lysed and the cDNA amplified using reverse transcription followed by PCR. The resulting cDNA quality is assessed on an agarose gel such as the one shown. After the DNA ladder in the first lane, the lanes show cDNA smears from single cells (indicated by the red brackets) alternating with amplification products from media controls. Smears of this quality (red brackets) are hybridized to Affymetrix microarrays.
<img alt="Figure 2" src="/files/ftp_upload/3824/3824fig2.jpg" /> 
Figure 2. Photos of the needles and aspirator tube assembly. Single cells are isolated using the capillary action of a pulled glass micropipette (inner diameter 0.5 mm, outer diameter 1.04 mm) and are transferred by the pressure of blowing into the aspirator. It is important that the aspirator tube is both long enough to easily manipulate and short enough to reliably discharge individual cells. A close-up view of the capillary tube pulled into a needle is shown in B.
<img alt="Figure 3" src="/files/ftp_upload/3824/3824fig3.jpg" /> 
Figure 3. Representative example of single cell cDNA smears and gene specific PCRs. To determine the quality of the single cell cDNA after amplification, 10 &mu;l of each single cell sample were run on a 1% agarose gel along with a negative control (A). A smear should appear between 500-2000 bp. Additional PCR-based screening for specific genes can help to identify/confirm the type of cell that was isolated and the amount of contamination present in the sample (B). Primers specific for the retinal ganglion cell markers Brn3b and Pax6 were tested to confirm the identity of these cells (Rows 1 and 2). To assess the amount of photoreceptor contamination in the preparation, primers specific for the photoreceptor marker Crx were used (Row 3). Subsets of ganglion cells may also be identified through screening for markers such as Tachykinin1 (Row 4).
<img alt="Figure 4" src="/files/ftp_upload/3824/3824fig4.jpg" /> 
Figure 4. Single cell expression of marker genes. The microarray results for selected genes expressed in 22 distinct single cells are shown in a heatmap format. The intensities from the microarray signals have been scaled to correspond with the intensity of the red color. Black indicates the absence of signal on the microarray. The retinal ganglion cell (RGC) precursors shown were isolated from embryonic time points, while the other cells were isolated from adult retinas.

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No conflicts of interest declared.

An ever-expanding number of studies are revealing robust cell-to-cell variability in populations that were believed to be more homogeneous with regard to their gene expression6,8. In at least one instance, this gene expression "noise" has been shown to play an important biological function13. Gene expression differences between individual cells are obscured using traditional whole-tissue methods. These experiments generate the expression profile of an "average" cell, which may not be representative14. Here we present a method for the isolation and identification of the gene expression patterns from single retinal cells. The study of individual cells allows researchers to probe the cellular heterogeneity underlying complex tissues, which is critical for determining distinctive gene expression patterns of various cell types. Additionally, single-cell isolation can provide insight into the gene programs driving the activity of individual cells in time points throughout development. Though especially useful in studying the nervous system, where the functioning of complex cellular networks can depend on the unique gene expression of rare cell types, this protocol can be adapted to isolate individual cells from various tissues.
In the early-developing murine retina, ganglion cells, horizontal cells, cone photoreceptor cells and amacrine cells are all generated in an overlapping window of time15. Additionally, each cell that has exited the cell cycle is in a different stage of maturation and this fact only adds to the complexity of the developing retina. Finally, for some types of retinal neurons, there exist numerous different morphologies (up to ~30 in the case of amacrine cells) in the mature retina16. Since the retina is such a mixture of cell types, microarray and serial analysis of gene expression (SAGE) based studies17-19 that relied on the homogenization of the entire retina are unable to uncover markers genes expressed in rare subtypes and unable to resolve dynamic gene expression differences between cell types, especially during development. To begin to understand the complexity of different cell types both in the adult retina and during retinal development, the single cell gene expression profiles of ~200 individual cells from many different time points have been analyzed2-5,8. The resulting data has provided a host of new markers for individual cell types and provided a window into important transitions in developmental time that were previously unappreciated.
The single cell expression profiles have revealed considerable heterogeneity in gene expression in amacrine cells4, where it was expected, and in M&uuml;ller glia cells, where it was unexpected5. While an examination of the single amacrine cell data revealed more than 450 genes that were expressed in amacrine cells and excluded from other retinal cell types, none of these genes were expressed in all the amacrine cells4. These results most likely arise from the fact that the amacrine cell class is extremely diverse and the underlying heterogeneity is a reflection of the distinct functions of these cells. These findings would not have been possible using whole tissue-based approaches. Finally, cycling retinal progenitor cells (RPCs) displayed the highest degree of heterogeneity in gene expression of any of the retinal cell types profiled8. Transcription factors were the class of genes with the highest degree of heterogeneity among the progenitor cells, suggesting that dynamic gene expression patterns could play a major role in the development of the different retinal cell types8. Again, these results would not have been possible without the use of the single cell gene expression profiling technique.
Single cell transcriptome studies can also be utilized to gain insight into disease mechanisms. In many neurodegenerative diseases, including retinal degenerative diseases, some neurons die while the neighboring neurons survive20. Understanding why some cells undergo apoptosis requires the identification of genes or gene programs that are altered in individual cells in these disease models. Whole-tissue models will again potentially obscure the changes since cells often do not die at the same time and, therefore, at any given time the tissue is a mixture of dying and surviving cells. Additionally, for many cancers the cell of origin is an open question. This is particularly the case with the childhood eye tumor retinoblastoma. Recently using the single cell profiling protocol detailed here, it was shown that individual cells from retinoblastomas possess gene expression programs of multiple cell types21. It appears that these cells are hybrids of undifferentiated progenitor cells and neurons21. These experiments required an analysis at the single cell level and would not have been possible with whole tissue approaches.
Alternative approaches
Linear amplification is an alternative to the PCR-based amplification protocol detailed here. While PCR-based amplification is believed to result in a skewing of abundance relationships, linear amplification is thought to maintain these relationships22. The technique of linear amplification has been used to profile several neuronal types23. However, direct comparisons between the two methods have indicated that the linear amplification technique can be associated with a high false negative rate24,25. We favor the PCR-based method detailed in this report for two reasons. First, in our experiments we focus on genes with robust expression differences between cells. Second, we have found a very good correlation between our single cell microarray profiling experiments and in situ hybridization studies for the same genes. In fact, to date we have performed in situ hybridizations for hundreds of genes and have observed at least 75% match the predicted pattern from the microarrays. This is most likely a conservative estimate since the most common reason for a discrepancy is a lack of signal from the in situ probe.

Reaction mixtures:
Cell Lysis buffer
<table border="0">
	<tbody>
		<tr>
			<td>0.45 &mu;</td>
			<td>10X reaction buffer (100 mM Tris-HCl pH 8.3, 500 mM KCl, 2 mM MgCl2)</td>
		</tr>
		<tr>
			<td>0.23 &mu;l</td>
			<td>10% NP-40</td>
		</tr>
		<tr>
			<td>0.23 &mu;l</td>
			<td>0.1M DTT</td>
		</tr>
		<tr>
			<td>0.05 &mu;l</td>
			<td>RNase Inhibitor (40 U/&mu;l)</td>
		</tr>
		<tr>
			<td>0.05 &mu;l</td>
			<td>SUPERase-In (20U/&mu;l)</td>
		</tr>
		<tr>
			<td>0.13 &mu;l</td>
			<td>Modified Oligo d(T) primer 
			(TATAGAATTCGCGGCCGCTCGCGATTTTTTTTTTTTTTTTTTTTTTTT)</td>
		</tr>
		<tr>
			<td>0.09 &mu;l</td>
			<td>dNTPs (2.5 mM each)</td>
		</tr>
		<tr>
			<td>3.27 &mu;l</td>
			<td>dH20 (use molecular biology grade for all reaction mixtures)</td>
		</tr>
	</tbody>
</table>
Tailing Reaction Mixture
<table border="0">
	<tbody>
		<tr>
			<td>0.18 &mu;l</td>
			<td>100 mM dATP</td>
		</tr>
		<tr>
			<td>0.6 &mu;l</td>
			<td>10X reaction buffer(100 mM Tris-HCl pH 8.3, 500 mM KCl, 2 mM MgCl2)</td>
		</tr>
		<tr>
			<td>4.62 &mu;l</td>
			<td>dH2O</td>
		</tr>
		<tr>
			<td>0.3 &mu;l</td>
			<td>TdT (400 U/&mu;l)</td>
		</tr>
		<tr>
			<td>0.3 &mu;l</td>
			<td>RNase H (2 U/&mu;l)</td>
		</tr>
	</tbody>
</table>
PCR Reaction Mixture
<table border="0">
	<tbody>
		<tr>
			<td>10 &mu;l</td>
			<td>10x Ex-Taq HS Buffer with Mg2+</td>
		</tr>
		<tr>
			<td>10 &mu;l</td>
			<td>2.5 mM dNTPs</td>
		</tr>
		<tr>
			<td>0.2 &mu;l</td>
			<td>Modified Oligo d(T) primer (10 &mu;g/&mu;l)</td>
		</tr>
		<tr>
			<td>1 &mu;l</td>
			<td>Ex-Taq HS Polymerase (5U/&mu;l)</td>
		</tr>
		<tr>
			<td>68.8 &mu;l</td>
			<td>dH2O</td>
		</tr>
	</tbody>
</table>
10X One-Phor All Buffer
<table border="0">
	<tbody>
		<tr>
			<td>0.5M</td>
			<td>Potassium acetate</td>
		</tr>
		<tr>
			<td>0.1M</td>
			<td>Tris acetate (pH 7.6)</td>
		</tr>
		<tr>
			<td>0.1M</td>
			<td>Magnesium acetate</td>
		</tr>
	</tbody>
</table>
Solutions:
10X Phosphate buffered saline (1 liter)
80g NaCl 
2g KCl 
14.4 g Na2PO4 
2.4 g KH2PO4 
adjust to pH 7.4 with NaOH and bring the volume to 1 liter with water
<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalog number</td>
			<td>Comments </td>
		</tr>
		<tr>
			<td>Vertical needle puller</td>
			<td>David Kopf Instruments</td>
			<td>Model 750</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Microcapillary tubes</td>
			<td>Sigma</td>
			<td>P0674</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Aspirator tube assembly</td>
			<td>Sigma</td>
			<td>A5177</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Corning filter tips (100-1000 &mu;l</td>
			<td>Fisher Scientific</td>
			<td>07-200-265</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hank's balanced salt solution (1X)</td>
			<td>Lonza BioWhittaker</td>
			<td>10-508F</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>HEPES buffer (1M)</td>
			<td>MP Biomedicals</td>
			<td>091688449</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Bovine serum albumin</td>
			<td>Sigma</td>
			<td>A9418</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>DNase I</td>
			<td>Roche</td>
			<td>04716728001</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>papain</td>
			<td>Worthington</td>
			<td>LS03126</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Superscript III</td>
			<td>Invitrogen</td>
			<td>18080-044</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>RNase Inhibitor</td>
			<td>Applied Biosystems</td>
			<td>AM2682</td>
			<td>These two RNase inhibitors seem to work the best. Contaminants present in other inhibitors can contribute to a background smear.</td>
		</tr>
		<tr>
			<td>SUPERase-In</td>
			<td>Applied Biosystems</td>
			<td>AM2694</td>
			<td>These two RNase inhibitors seem to work the best. Contaminants present in other inhibitors can contribute to a background smear.</td>
		</tr>
		<tr>
			<td>Modified Oligo d(T) primer (gel purified)</td>
			<td>Oligos ETC</td>
			<td>&nbsp;</td>
			<td>We have used primers purchased from many different companies and have seen the most reliable and reproducible results from Oligos ETC.</td>
		</tr>
		<tr>
			<td>T4 gene 32 protein</td>
			<td>New England Biolabs</td>
			<td>M0300L</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Exonuclease I</td>
			<td>New England Biolabs</td>
			<td>M0293S</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>TdT</td>
			<td>Roche</td>
			<td>3 333 574</td>
			<td>As with other reagents, using a TdT enzyme from other companies gave more variable results.</td>
		</tr>
		<tr>
			<td>RNase H</td>
			<td>Invitrogen</td>
			<td>18021-014</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Ex-Taq HS Polymerase</td>
			<td>Takara</td>
			<td>RR006A</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Biotin N6-ddATP</td>
			<td>Enzo Biosciences</td>
			<td>42809</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>
Table 1. Specific reagents and equipment.

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.

Watch this Scientific Journal Video about Single-cell Profiling of Developing and Mature Retinal Neurons at JoVE.com

Single-cell Profiling of Developing and Mature Retinal Neurons

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 ...

single-cell-profiling-of-developing-and-mature-retinal-neurons

Nanyang Technological University

Research

JoVE Journal

Neuroscience

14.1K Views.  Iowa State University. 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.

Video: Single-cell Profiling of Developing and Mature Retinal Neurons

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 ...

Profiling Voltage-gated Potassium Channel mRNA Expression in Nigral Neurons using Single-cell RT-PCR Techniques

In mammalian central nervous system, different types of neurons with diverse molecular and functional characteristics are intermingled with each other, difficult to separate and also not easily identified by their morphology. Thus, it is often difficult to analyze gene expression in a specific neuron type. Here we document a procedure that combines whole-cell patch clamp recording techniques with single-cell reverse transcription polymerase chain reaction (scRT-PCR) to profile mRNA expression in different types of neurons in the substantial nigra. Electrophysiological techniques are first used to record the neurophysiological and functional properties of individual neurons. Then, the cytoplasm of single electrophysiologically characterized nigral neurons is aspirated and subjected to scRT-PCR analysis to obtain mRNA expression profiles for neurotransmitter synthesis enzymes, receptors, and ion channels. The high selectivity and sensitivity make this method particularly useful when immunohistochemistry can not be used due to a lack of suitable antibody or low expression level of the protein. This method is also applicable to neurons in other brain areas.

Neurons are first characterized electrophysiologically. Then the cytoplasm from the recorded neuron is aspirated and subjected to reverse transcription-PCR analysis to detect the expression of mRNAs for neurotransmitter synthesis enzymes, ion channels, and receptors.

In mammalian central nervous system, different types of neurons with diverse molecular and functional characteristics are intermingled with each other, ...

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.

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 ...

Simultaneous Whole-cell Recordings from Photoreceptors and Second-order Neurons in an Amphibian Retinal Slice Preparation

One of the central tasks in retinal neuroscience is to understand the circuitry of retinal neurons and how those connections are responsible for shaping the signals transmitted to the brain. Photons are detected in the retina by rod and cone photoreceptors, which convert that energy into an electrical signal, transmitting it to other retinal neurons, where it is processed and communicated to central targets in the brain via the optic nerve. Important early insights into retinal circuitry and visual processing came from the histological studies of Cajal1,2 and, later, from electrophysiological recordings of the spiking activity of retinal ganglion cells - the output cells of the retina3,4.
A detailed understanding of visual processing in the retina requires an understanding of the signaling at each step in the pathway from photoreceptor to retinal ganglion cell. However, many retinal cell types are buried deep in the tissue and therefore relatively inaccessible for electrophysiological recording. This limitation can be overcome by working with vertical slices, in which cells residing within each of the retinal layers are clearly visible and accessible for electrophysiological recording.
Here, we describe a method for making vertical sections of retinas from larval tiger salamanders (Ambystoma tigrinum). While this preparation was originally developed for recordings with sharp microelectrodes5,6, we describe a method for dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells in which we manipulate the photoreceptor's membrane potential while simultaneously recording post-synaptic responses in horizontal or bipolar cells. The photoreceptors of the tiger salamander are considerably larger than those of mammalian species, making this an ideal preparation in which to undertake this technically challenging experimental approach. These experiments are described with an eye toward probing the signaling properties of the synaptic ribbon - a specialized synaptic structure found in a only a handful of neurons, including rod and cone photoreceptors, that is well suited for maintaining a high rate of tonic neurotransmitter release7,8 - and how it contributes to the unique signaling properties of this first retinal synapse.

We describe the preparation of thin retinal slices from aquatic tiger salamanders (Ambystoma tigrinum) and explain how we use these slices to study synaptic processing in the retina by obtaining dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells.

One of the central tasks in retinal neuroscience  is to understand the circuitry of retinal neurons and how those connections are  responsible for shaping ...

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 ...

Isolation of Adult Spinal Cord Nuclei for Massively Parallel Single-nucleus RNA Sequencing

Probing an individual cell&#39;s gene expression enables the identification of cell type and cell state. Single-cell RNA sequencing has emerged as a powerful tool for studying transcriptional profiles of cells, particularly in heterogeneous tissues such as the central nervous system. However, dissociation methods required for single cell sequencing can lead to experimental changes in the gene expression and cell death. Furthermore, these methods are generally restricted to fresh tissue, thus limiting studies on archival and bio-bank material. Single nucleus RNA sequencing (snRNA-Seq) is an appealing alternative for transcriptional studies, given that it accurately identifies cell types, permits the study of tissue that is frozen or difficult to dissociate, and reduces dissociation-induced transcription. Here, we present a high-throughput protocol for rapid isolation of nuclei for downstream snRNA-Seq. This method enables isolation of nuclei from fresh or frozen spinal cord samples and can be combined with two massively parallel droplet encapsulation platforms.

Here, we present a protocol to rapidly isolate high-quality nuclei from the fresh or frozen tissue for downstream massively parallel RNA sequencing. We include detergent-mechanical and hypotonic-mechanical tissue disruption and cell lysis options, both of which can be used for isolation of nuclei.

Probing an individual cell&#39;s gene expression enables the identification of cell type and cell state. Single-cell RNA sequencing has emerged as a ...

Single-cell RNA Sequencing of Fluorescently Labeled Mouse Neurons Using Manual Sorting and Double In Vitro Transcription with Absolute Counts Sequencing (DIVA-Seq)

Single-cell RNA sequencing (RNA-seq) is now a widely implemented tool for assaying gene expression. Commercially available single-cell RNA-sequencing platforms process all input cells indiscriminately. Sometimes, fluorescence-activated cell sorting (FACS) is used upstream to isolate a specifically labeled population of interest. A limitation of FACS is the need for high numbers of input cells with significantly labeled fractions, which is impractical for collecting and profiling rare or sparsely labeled neuron populations from the mouse brain. Here, we describe a method for manually collecting sparse fluorescently labeled single neurons from freshly dissociated mouse brain tissue. This process allows for capturing single-labeled neurons with high purity and subsequent integration with an in vitro transcription-based amplification protocol that preserves endogenous transcript ratios. We describe a double linear amplification method that uses unique molecule identifiers (UMIs) to generate individual mRNA counts. Two rounds of amplification results in a high degree of gene detection per single cell.

Single-cell RNA Sequencing of Fluorescently Labeled Mouse Neurons Using Manual Sorting and Double In Vitro Transcription with Absolute Counts Sequencing DIVA-Seq

This protocol describes the manual sorting procedure to isolate single fluorescently labeled neurons followed by in vitro transcription-based mRNA amplification for high-depth single-cell RNA sequencing.

Single-cell RNA sequencing (RNA-seq) is now a widely implemented tool for assaying gene expression. Commercially available single-cell RNA-sequencing ...

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.

Preparation of high-quality mouse eye sections for immunohistochemistry (IHC) is critical for assessing the retinal structure and function and for ...

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding environment and initiate contacts with potential synaptic partners. Although the connection between dendritic branch dynamics and synaptogenesis is well established, how developmental and activity-dependent processes regulate dendritic branch dynamics is not well understood. This is partly due to the technical difficulties associated with the live imaging and quantitative analyses of these fine structures using an in vivo system. We established a method to study dendrite dynamics using Drosophila larval ventral lateral neurons (LNvs), which can be individually labeled using genetic approaches and are accessible for live imaging. Taking advantage of this system, we developed protocols to capture branch dynamics of the whole dendritic arbor of a single labeled LNv through time-lapse live imaging. We then performed post-processing to improve image quality through drift correction and deconvolution, followed by analyzing branch dynamics at the single-branch level by annotating spatial positions of all branch terminals. Lastly, we developed R scripts (Supplementary File) and specific parameters to quantify branch dynamics using the coordinate information generated by the terminal tracing. Collectively, this protocol allows us to achieve a detailed quantitative description of branch dynamics of the neuronal dendritic arbor with high temporal and spatial resolution. The methods we developed are generally applicable to sparsely labeled neurons in both in vitro and in vivo conditions.

Time-lapse Live Imaging and Quantification of Fast Dendritic Branch Dynamics in Developing Drosophila Neurons

Here, we describe the method we employed to image highly motile dendritic filopodia in a live preparation of the Drosophila larval brain, and the protocol we developed to quantify time-lapse 3D imaging datasets for quantitative assessments of dendrite dynamics in developing neurons.

Highly motile dendritic filopodia are widely present in neurons at early developmental stages. These exploratory dynamic branches sample the surrounding ...

Single-Cell Electroporation across Different Organotypic Slice Culture of Mouse Hippocampal Excitatory and Class-Specific Inhibitory Neurons

Electroporation has established itself as a critical method for transferring specific genes into cells to understand their function. Here, we describe a single-cell electroporation technique that maximizes the efficiency (~80%) of in vitro gene transfection in excitatory and class-specific inhibitory neurons in mouse organotypic hippocampal slice culture. Using large glass electrodes, tetrodotoxin-containing artificial cerebrospinal fluid and mild electrical pulses, we delivered a gene of interest into cultured hippocampal CA1 pyramidal neurons and inhibitory interneurons. Moreover, electroporation could be carried out in cultured hippocampal slices up to 21 days in vitro with no reduction in transfection efficiency, allowing for the study of varying slice culture developmental stages. With interest growing in examining the molecular functions of genes across a diverse range of cell types, our method demonstrates a reliable and straightforward approach to in vitro gene transfection in mouse brain tissue that can be performed with existing electrophysiology equipment and techniques.

Presented here is a protocol for single-cell electroporation that can deliver genes in both excitatory and inhibitory neurons across a range of in vitro hippocampal slice culture ages. Our approach provides precise and efficient expression of genes in individual cells, which can be used to examine cell-autonomous and intercellular functions.

Electroporation has established itself as a critical method for transferring specific genes into cells to understand their function. Here, we describe a ...