This work was supported by NIH grant R01-EY020535 (B.T.S), International Retinal Research Foundation and Karl Kirchgessner Foundation (B.T.S).

In all experimental procedures, animals were treated according to the regulations in the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and in compliance with protocols approved by Weill Cornell Medical College. Animals were euthanized by carbon dioxide and subsequent cervical dislocation.
1. Dissection of Wholemount Retina
<ol>
	<li>After the mouse is euthanized, enucleate its eyes and place them into oxygenated HEPES-buffered Ringer&#39;s solution (Table 1).</li>
	<li>Remove the cornea by cutting in a circular path along the ora serrata with small scissors, holding the eye at the limbus with forceps. Pull out the lens with forceps.</li>
	<li>Remove as much vitreous as possible, using blunted-edge forceps. This step is important for bolus injections and electrophysiological recordings; it is not necessary for when performing immunohistochemistry alone.</li>
	<li>Detach the retina from the eyecup by positioning forceps between the retina and the eyecup, moving the forceps slowly around the circumference. Stabilize the eye cup by holding the sclera firmly with a second set of forceps. Cut the optic nerve at the location between the retina and the eyecup to ease the separation of the retina from the eyecup.</li>
	<li>The retina can be dissected into pieces (e.g. quarters) or used as a whole. To cut into pieces, use scissors to make incisions from the retinal periphery all the way down to the optic nerve. Alternatively, to use the entire retina, make incisions from the periphery halfway to the optic nerve. The incisions will help to make a flat preparation.</li>
</ol>
2. Mounting onto Hydrophilized PTFE Membrane Insert
<ol>
	<li>Prepare the membrane as described below. When physiological recordings or dye injection experiments are performed, shorten the holder by using either a lathe or a handsaw so that the remaining ring has a height of 1 mm. 
	Note: The culture plate inserts may be either unmodified or modified (Figure 1A). The choice of the configuration is dictated by the purpose of the experiment. For immunostaining, an unmodified insert is suitable.</li>
	<li>Polish down rough surfaces with fine (300 grit or higher) sandpaper.</li>
	<li>(Optional) Remove the feet underneath the ring with scissors.</li>
	<li>To remove any debris, rinse or sonicate the membrane in distilled water for 2-3 min.</li>
	<li>Mount the tissue onto the membrane (Figure 1A). Place the tissue with some solution on top of the membrane with a plastic transfer pipette. Cut off the narrow tip of the pipette if necessary, to avoid having the tissue sample adhere to the pipette during transfer.</li>
	<li>Attach the retina with ganglion cells (GCs) up: on the membrane, gradually remove solution while spreading the retina and unfolding the folds with forceps.</li>
	<li>On the opposite side of the membrane, position an empty syringe (1 ml) directly underneath the tissue. Apply suction by pulling back on the plunger to a volume of 0.2-0.3 ml. Increase the surface area of suction by cutting the holding tip of the syringe prior to use. The tissue does not require further treatment for adhesion, as it is physically enmeshed within the membrane.</li>
</ol>
3. Bolus Injection of Isolectin Alexa 488
<ol>
	<li>Prepare patch pipettes and staining solution. Pull patch pipettes from borosilicate capillary glass with a Flaming/Brown puller, with a resistance of ~1 M&#8486;. Gently break the tip of the pipette.</li>
	<li>Make staining solution by adding 0.3 &#956;l of isolectin Alexa 488 stock solution to 10 &#956;l of the HEPES-buffered Ringer&#39;s solution described in step 1.2.</li>
	<li>Transfer sample to the patch-clamp setup for injection (Figure 1C). Place the membrane with attached tissue into the recording chamber perfused with warmed (37 &#176;C) and oxygenated HEPES-buffered Ringer&#39;s solution.</li>
	<li>Place the pipette containing dye against the inner limiting membrane (ILM, Figure 2A). Using micromanipulator, further advance the pipette against the ILM until it penetrates it. Inject the staining solution with a pressure 10-20 psi for 1 sec using a picospritzer. Single or multiple locations may be targeted for injection (Figure 2B). If a picospritzer is not available, a 10 ml syringe can be used to deliver a positive air pressure for injection.</li>
	<li>Incubate injected tissue for 10 min. During incubation, the injected tissue can be viewed under epifluorescence illumination.</li>
	<li>After labeling, the preparation may be taken to a microscope for imaging. Transfer the insert to a glass-bottom culture dish, placing retina side down (Figure 1C).</li>
	<li>Add a drop of HEPES solution to keep the specimen moist.</li>
	<li>Place 2-3 spots of vacuum grease around the perimeter of the plastic ring to firmly secure the insert to the bottom of the culture dish.</li>
	<li>Transfer the dish to the microscope stage.</li>
</ol>
4. Paraformaldehyde and Carbodiimide Fixation
<ol>
	<li>Fix the sample by transferring the insert into fixative for 15 min. Longer or shorter fixation times may be used, dependent on application. Both fixatives can be effectively used to fix GFP, isolectin, or fluorescent Alexa hydrazides (e.g. used for filling the cells in patch-clamp recordings). 
	Note: For fixation, either 4% carbodiimide or 4% paraformaldehyde solutions may be used. To maintain adherence, fixation should be done after the tissue has already been attached to the membrane. The tissue can be fixed immediately after mounting, or after physiological recording and/or bolus injection.</li>
	<li>Rinse the tissue in phosphate buffer (PB) for 30 min. After this, the tissue may be processed either as a wholemount (step 5) or cut into vertical sections and then processed (steps 6 and 7).</li>
</ol>
5. Immunostaining of Retinal Wholemounts
<ol>
	<li>In a 24-well dish (Figure 1D) block retinal pieces on the membrane for 1 hr in a staining solution.</li>
	<li>Dilute mouse anti-PSD-95 primary antibodies 1:300 in the staining solution and apply for 48 hr at room temperature in dark. Add ~100-150 &#956;l antibody solution in each well with the tissue. To prevent drying out, fill unused wells with distilled water.</li>
	<li>Rinse for 20 min in PB, 3x.</li>
	<li>Incubate for 24 hr in donkey anti-mouse Alexa 568 secondary antibody dissolved 1:500 in the staining solution.</li>
	<li>Rinse for 20 min in PB, 3x.</li>
	<li>Place the insert with the stained tissue into a standard 3 cm Petri dish filled with PB solution. With small scissors, cut out the portion of the membrane containing the retina from the ring.</li>
	<li>Transfer the retina on the membrane onto a slide retina up. Remove PB, add mounting medium. Place small pieces of a broken cover slip glass in between the slide and the coverslip glasses to avoid extensive squeezing and damage to the retina. Place coverslip.</li>
	<li>Seal the coverslip in place with nail polish. Keep slides in the dark, in a refrigerator.</li>
</ol>
6. Vertical Sectioning of Membrane-mounted Tissue
<ol>
	<li>After fixation, submerse the tissue on the membrane into the sucrose gradient solutions of 10%, 20%, and 30% sucrose for 30 min, 1 hr, and overnight, respectively. The overnight step is performed at 4 &#176;C in refrigerator, all other steps at room temperature.</li>
	<li>Using scissors, remove the membrane with the tissue from the plastic insert ring.</li>
	<li>Place the cut off membrane onto Parafilm with retina up and remove as much solution as possible with Kimwipes.</li>
	<li>Put the Parafilm with the tissue into the cryostat chamber and add a drop of OCT medium. Use the quick freezer device to accelerate the freezing.</li>
	<li>Detach the frozen sample and flip it over.</li>
	<li>Cut excess frozen OCT. Leave approximately 2 mm around the sample on three sides, and leave 5-6 mm from the side which will be attached to the holder.</li>
	<li>Cover exposed retinal side (membrane side) with a drop of OCT. Leave the attaching area free.</li>
	<li>Freeze the membrane with retinal tissue. Now the sample is ready for placing onto a cutting holder.</li>
	<li>Place a drop of OCT onto a cold holder ring. Immediately insert the frozen sample into the drop.</li>
	<li>Cut sections 10-20 &#956;m thick, setting cryostat temperature at -19 &#176;C. Use warm (room temperature) polysine adhesion slides. Leave the slides with the attached sections onto a heating platform for ~2-3 min (41 &#176;C) in dark to ensure firm adhesion of sections.</li>
	<li>Put the slides into a storage box and keep at -20 &#176;C until use.</li>
</ol>
7. Immunostaining of Retinal Sections
<ol>
	<li>Take the slides with the sections out of the freezer. Let them stay at room temperature for 1 min to remove the condensate.</li>
	<li>Encircle the sections with a liquid blocker to create a barrier for further incubation.</li>
	<li>Place the slides into the incubation box. Cover the bottom of the box with water to prevent evaporation of the antibody solution.</li>
	<li>Rinse with PB for 5 min.</li>
	<li>Prepare the same staining solutions and follow the steps as in step 5. The incubation with the primary antibody is overnight and with the secondary 1 hr. Do not use broken glass between the coverslip and the slide.</li>
</ol>

<ol>
	<li>Csaba, G., Kovacs, P., Pallinger, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EDAC+fixation+increases+the+demonstrability+of+biogenic+amines+in+the+unicellular+Tetrahymena:+a+flow+cytometric+and+confocal+microscopic+comparative+analysis.">EDAC fixation increases the demonstrability of biogenic amines in the unicellular Tetrahymena: a flow cytometric and confocal microscopic comparative analysis.</a> Cell Biol. Int. 30, 345-348 (2006).</li><li>Fletcher, E. L., Hack, I., Brandstatter, J. H., Wassle, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+localization+of+NMDA+receptor+subunits+in+the+rat+retina.">Synaptic localization of NMDA receptor subunits in the rat retina.</a> J. Comp. Neurol. 420, 98-112 (2000).</li><li>Gastinger, M. J., O'Brien, J. J., Larsen, N. B., Marshak, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Histamine+immunoreactive+axons+in+the+macaque+retina.">Histamine immunoreactive axons in the macaque retina.</a> Invest. Ophthalmol. Vis. Sci. 40, 487-495 (1999).</li><li>Haverkamp, S., Wassle, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunocytochemical+analysis+of+the+mouse+retina.">Immunocytochemical analysis of the mouse retina.</a> J. Comp. Neurol. 424, 1-23 (2000).</li><li>Koulen, P., Fletcher, E. L., Craven, S. E., Bredt, D. S., Wassle, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunocytochemical+localization+of+the+postsynaptic+density+protein+PSD-95+in+the+mammalian+retina.">Immunocytochemical localization of the postsynaptic density protein PSD-95 in the mammalian retina.</a> J. Neurosci. 18, 10136-10149 (1998).</li><li>Pena, J. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=miRNA+in+situ+hybridization+in+formaldehyde+and+EDC-fixed+tissues.">miRNA in situ hybridization in formaldehyde and EDC-fixed tissues.</a> Nat. Methods. 6, 139-141 (2009).</li><li>Tymianski, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+use+for+a+carbodiimide+compound+for+the+fixation+of+fluorescent+and+non-fluorescent+calcium+indicators+in+situ+following+physiological+experiments.">A novel use for a carbodiimide compound for the fixation of fluorescent and non-fluorescent calcium indicators in situ following physiological experiments.</a> Cell Calcium. 21, 175-183 (1997).</li><li>Sagdullaev, B. T., Eggers, E. D., Purgert, R., Lukasiewicz, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonlinear+interactions+between+excitatory+and+inhibitory+retinal+synapses+control+visual+output.">Nonlinear interactions between excitatory and inhibitory retinal synapses control visual output.</a> J. Neurosci. 31, 15102-15112 (2011).</li><li>Toychiev, A. H., Sagdullaev, B., Yee, C. W., Ivanova, E., Sagdullaev, B. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+time+and+cost+efficient+approach+to+functional+and+structural+assessment+of+living+neuronal+tissue.">A time and cost efficient approach to functional and structural assessment of living neuronal tissue.</a> J. Neurosci. Methods. 214, 105-112 (2013).</li><li>Yee, C. W., Toychiev, A. H., Sagdullaev, B. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Network+deficiency+exacerbates+impairment+in+a+mouse+model+of+retinal+degeneration.">Network deficiency exacerbates impairment in a mouse model of retinal degeneration.</a> Front Syst. Neurosci. 6, 8 (2012).</li><li>Ivanova, E., Muller, U., Wassle, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+glycinergic+input+to+bipolar+cells+of+the+mouse+retina.">Characterization of the glycinergic input to bipolar cells of the mouse retina.</a> Eur. J. Neurosci. 23, 350-364 (2006).</li><li>Dmitrieva, N. A., Strang, C. E., Keyser, K. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+alpha+7+nicotinic+acetylcholine+receptors+by+bipolar,+amacrine,+and+ganglion+cells+of+the+rabbit+retina.">Expression of alpha 7 nicotinic acetylcholine receptors by bipolar, amacrine, and ganglion cells of the rabbit retina.</a> J. Histochem. Cytochem. 55, 461-476 (2007).</li></ol>

The authors have no competing financial interests.

The hydrophilized PTFE membrane&#39;s high biocompatibility and transparency in solutions is advantageous when working with living tissue. This preparation has been successfully used in earlier work for patch-clamp recordings of light responses8-10. Here, we show how this approach can be adopted for structural analysis of the wholemount retina.
The bolus loading technique can be effectively used for labeling living tissue. This technique is aided by the structural support of the membrane insert. Applications include loading of calcium indicator9, staining of the tissue with the fluorescent markers11, or labeled ligands12.
Fixed tissue processing can also benefit from the use of a supporting membrane. Differences in the appearance of postsynaptic density proteins depend on fixation time and the nature of the fixative5. Longer fixation times lead to excellent tissue preservation but poor antibody recognition of synaptic proteins that are cross-linked by the fixative. Shorter fixation times result in poor tissue preservation but bright punctate labeling. Similar results have been obtained for many other synaptic structural proteins and ion channels by different groups1-7. Using the membrane for structural support enables the use of carbodiimide to simultaneously achieve short fixation times, well-defined bright labeling of synaptic structures and adequate overall tissue quality for both wholemount and cryostat section preparations. The preparations are stable and easy to handle, with reproducible results.
Several critical steps should be carefully performed to ensure high quality and reproducible results. For bolus injections and electrophysiological recordings, it is important to remove as much of the vitreous as possible when dissecting the retina. The absence of the vitreous will enable easy access of the patch pipette to the cells. The vitreous does not interfere with the described immunohistochemistry techniques. To maintain firm adherence, fixation should be done after the tissue has already been attached to the membrane. For cryostat sectioning, the sucrose solutions must be completely removed prior to adding the embedding OCT medium. Failure to remove the liquid may result in splitting the cut section along the frozen tissue and damaging it. The slides with the attached sections should be kept for 2-3 min on a heating platform. If this step is not performed, the tissue may detach from the slide during the staining procedure.
There are several limitations of this technique. First, once the tissue adheres to the membrane, it cannot be removed without incurring damage, as the tissue is enmeshed within the membrane. Second, tissue cannot be attached to the membrane after it has been fixed. Third, while the membrane is fully transparent in water-based mediums, the membrane may look slightly opaque when in mounting medium, compromising light transmission. This is mitigated by sectioning, since this removes the membrane from the path of light transmitted to the tissue.

Performing immunohistochemistry in delicate intact tissues runs the risk of damage during handling and transfer. This can occur either in brain slices or other thin tissue, such as degenerated retina. Additionally, there are certain methods of tissue fixation that can be advantageous for immunostaining of neuronal structures, but result in compromised structural stability, precluding their use. A particular example of this is carbodiimide-based fixation, which is superior for staining receptors and hormones1-7 but is commonly avoided due to the instability of the fixed tissue.
Here, we describe a procedure which utilizes a hydrophilized PTFE membrane to structurally support delicate tissue, either fixed or unfixed, for a variety of staining techniques. The supporting membrane allows for the manipulation of delicate tissue before and after fixation, allowing for several steps of processing while minimizing the risk of tissue damage. Overall, this simple method to preserve tissue integrity provides the opportunity to use techniques that might otherwise be avoided. As such, this approach could also be successfully used for preparation of wide variety of tissues such as brain slices that become fragile following slicing procedures.

<table border="0" cellpadding="0" cellspacing="0" width="625">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="44">
			<td height="44" style="height:44px;width:260px;">Millicell Cell Culture Insert, 12 mm, 
			hydrophilic PTFE (Biopore), 0.4 &#956;m</td>
			<td style="width:183px;">Millipore</td>
			<td style="width:108px;">PICM01250</td>
			<td style="width:75px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Insulin syringe, 1 ml</td>
			<td>Beckton Dickinson</td>
			<td>309659</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Scissors</td>
			<td>Fine Science Tools</td>
			<td>15003-08</td>
			<td>dissection</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Forceps, Dumont #55, inox</td>
			<td>Fine Science Tools</td>
			<td>11255-20</td>
			<td>dissection</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cryostat</td>
			<td>Leica</td>
			<td>various</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Confocal Microscope System</td>
			<td>Nikon</td>
			<td>various</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Capillary glass</td>
			<td>World Precision Instruments</td>
			<td>1B150F-4</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">P-97 Flaming/Brown Micropipette Puller</td>
			<td>Sutter Instrument Co.</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Picospritzer III</td>
			<td>Parker Hannifin</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Glass Bottom Culture Dishes</td>
			<td>MatTek Corporation</td>
			<td>P35G-0-14-C</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Petri dish</td>
			<td>Falcon</td>
			<td>1008</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Disposable Graduated Transfer Pipettes</td>
			<td>VWR</td>
			<td>16001-180</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Multiwell plates, 24-well</td>
			<td>Beckton Dickinson</td>
			<td>351147</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cover glass, #1</td>
			<td>Electron Microscopy Sciences</td>
			<td>72200-30</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Polysine adhesion slides</td>
			<td>Electron Microscopy Sciences</td>
			<td>63412-01</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Microscope slides</td>
			<td>Globe</td>
			<td>1301</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Liquid Blocker</td>
			<td>Electron Microscopy Sciences</td>
			<td>71312</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">VECTASHIELD mounting medium</td>
			<td>Vector Laboratories</td>
			<td>H-1000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">OCT medium</td>
			<td>Sakura</td>
			<td>4583</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Parafilm laboratory film</td>
			<td>Fisher</td>
			<td>13-374-10</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dow Corning high vacuum grease</td>
			<td>Sigma-Aldrich</td>
			<td>Z273554</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Mouse anti-PSD95</td>
			<td>Millipore</td>
			<td>MABN68</td>
			<td>antibody</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Donkey anti-mouse Alexa 568</td>
			<td>Invitrogen</td>
			<td>A10037</td>
			<td>antibody</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Isolectin Alexa 488</td>
			<td>Invitrogen</td>
			<td>I21411</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Chemiblocker</td>
			<td>Chemicon</td>
			<td>2170S</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>T9284</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sulforhodamine</td>
			<td>Sigma-Aldrich</td>
			<td>341738</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Carbodiimide</td>
			<td>Thermo Scientific</td>
			<td>22980</td>
			<td>EDC</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Paraformaldehyde</td>
			<td>Sigma-Aldrich</td>
			<td>P6148</td>
			<td></td>
		</tr>
	</tbody>
</table>

optimized protocol for retinal wholemount preparation for imaging and immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

Demonstrated here are two representative experiments that benefit from the use of a hydrophilized PTFE membrane to support wholemount retina during staining procedures. The first experiment, illustrates the method for fast and simple characterization of retinal vasculature, an elaborate network of blood vessels that span across multiple retinal layers (Figure 2). This approach combines bolus loading of isolectin with immersion labeling with SRH. This is an efficient method of labeling living tissue that can be visualized and scanned almost immediately. In the retina, blood vessels can be labeled from superficial to the deep layer (Figure 2E). In contrast to relatively well diffusible SRH, isolectin does not permeate well across the inner limiting membrane (ILM). This limitation is overcome by injecting the solution containing isolectin underneath the ILM with a glass pipette. For SRH blood vessels labeling, it is better to stain the tissue before mounting on the insert. The SRH stains only living tissue and the labeling disappears following fixation with either paraformaldehyde or carbodiimide.
The second experiment (Figure 3), shows the advantages of fixation with carbodiimide over conventional fixation with paraformaldehyde when labeling synaptic proteins. In general, the use of carbodiimide fixative is avoided because it results in relatively fragile tissue. However, mounting the tissue on a membrane ensures sufficient structural support, mitigating this issue. With a short period of &#64257;xation with carbodiimide, followed by staining for the synaptic marker PSD-95 immuno&#64258;uorescence in the IPL had a bright punctate appearance, suggesting that individual synapses were distinguished (Figure 3B). In contrast, when using conventional paraformaldehyde fixative, identification of the synaptic components is less clear (Figure 3E). Similar results were obtained in the retinal cryostat sections (Figures 3C and F).
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51018/51018fig1.jpg" width="600px" /> 
Figure 1. Versatile use of hydrophilized PTFE membranes. (A) Living retinal wholemount attached to an unmodified 12 mm membrane insert (left) and a quarter of retina attached to an insert with the holder portion removed. See video demonstrating mounting procedure. (B) Living retina on a stage of an upright microscope within a patch-clamp setup. (C) Living retina on a stage of an inverted confocal microscope. (D) Free floating retinal tissue (arrow) and the tissue mounted on membrane insert (arrowhead). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51018/51018fig1highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51018/51018fig2.jpg" width="600px" /> 
Figure 2. Focal and global labeling of the vasculature in the living retinal wholemount after injection of isolectin Alexa 488. (A) DIC image of the retina with a pipette filled with isolectin above the inner limiting membrane (ILM) prior to injection. (B) The same area as in A after penetration of the pipette beneath the ILM and pressure injection of the pipette solution. The spot with the clearly visible ganglion cells indicate that the solution has been injected between the ILM and the ganglion cell layer. (C-H) Confocal images of a live retinal wholemount labeled with sulforhodamine (SRH, red) and isolectin (green). (C) Isolectin applied with multiple bolus injections (asterisks). The size of the labeled area varies according to the pressure of the injection. SRH labels the entire vasculature. (D) Projection of a z-stack under high magnification shows all layers of vasculature and brightly labeled microglia (arrow). (E) Z-stack from the area highlighted in D rotated 90&#176;. (F-H) A view of the individual layers of blood vessels corresponding to labels in E. OPL-outer plexiform layer, INL-inner nuclear layer, GCL-ganglion cell layer. Scale bars: 50 &#956;m for A-B and D-H, 1 mm for C. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51018/51018fig2highres.jpg" target="_blank">Click here to view larger image</a>.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51018/51018fig3.jpg" width="600px" /> 
Figure 3. Role of the fixative in the antibody labeling of synaptic structures. Retinal wholemounts fixed with either carbodiimide (A-C) or paraformaldehyde (D-F). (A,D) Ganglion cells expressing green fluorescent proteinin B6.Cg-Tg(Thy1-YFPH)2Jrs/J mice (Jackson Lab. Stock #0033548). Squares indicate areas of high magnification shown in B and E. (B,E) Staining for the synaptic marker PSD-95 (red). Projections are within a narrow focal plane (two confocal images spaced 0.3 &#956;m apart). (C,F) Single confocal image of vertical section stained for PSD-95. Note that the axon terminals of photoreceptors within the OPL are overexposed to show weaker staining for synapses in the IPL (left panels). Inserts (right panels) show high magnifications of the areas highlighted on the left panels. OPL-outer plexiform layer, INL-inner nuclear layer, IPL-inner plexiform layer, GCL-ganglion cell layer. Scale bars = 50 &#956;m in A and D; 10 &#956;m in B, C, E, and F. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51018/51018fig3highres.jpg" target="_blank">Click here to view larger image</a>.

Watch this Scientific Journal Video about Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry at JoVE.com

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

optimized-protocol-for-retinal-wholemount-preparation-for-imaging

Nanyang Technological University

Research

JoVE Journal

Biology

23.0K Aufrufe.  Weill Medical College of Cornell University. The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Serie: Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Neutrophil Isolation Protocol

Neutrophil polymorphonuclear granulocytes (PMN) are the most abundant leukocytes in humans and among the first cells to arrive on the site of inflammatory immune response. Due to their key role in inflammation, neutrophil functions such as locomotion, cytokine production, phagocytosis, and tumor cell combat are extensively studied. To characterize the specific functions of neutrophils, a clean, fast, and reliable method of separating them from other blood cells is desirable for in vitro studies, especially since neutrophils are short-lived and should be used within 2-4 hours of collection. Here, we demonstrate a standard density gradient separation method to isolate human neutrophils from whole blood using commercially available separation media that is a mixture of sodium metrizoate and Dextran 500. The procedure consists of layering whole blood over the density gradient medium, centrifugation, separation of neutrophil layer, and lysis of residual erythrocytes. Cells are then washed, counted, and resuspended in buffer to desired concentration. When performed correctly, this method has been shown to yield samples of >95% neutrophils with >95% viability.

Neutrophils are among the first cells to arrive on the site of inflammatory immune response, and their functions and mechanisms have been studied extensively in vitro. We demonstrate a standard density gradient separation method to isolate human neutrophils from whole blood using commercially available separation media.

Neutrophile Isolation Protocol

Neutrophil polymorphonuclear granulocytes (PMN) are the most abundant leukocytes in humans and among the first cells to arrive on the site of inflammatory ...

Forschung

Biologie

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Die Herstellung Drosophila Embryonen für Live-Imaging mit dem Hanging-Drop-Protokoll

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

In vivo-like Organotypic Murine Retinal Wholemount Culture

Targeted ablations of genes and analysis of animal models is the classical strategy for enrolling specific retinal gene function. However, transgenic, retina-specific or conditional knockout mouse models often display early lethality or suffer from severe malformations, preventing an analysis beyond embryonic or early postnatal stages.
Primary cell culture is an alternative to investigate the effects of exogenously applied recombinant factors, overexpression of genes or siRNA-mediated gene knockdown in a controlled environment. Dissociated cell culture has the advantage that the endogenous signals reaching the target cells are reduced, thereby facilitating the identification of exogenously triggered effects after pharmacological manipulation. However, important cell-cell interactions are initially destroyed by enzymatic digestion or mechanical dissociation, even if re-aggregated retinospheroid cultures1 are used.
By contrast, organotypic retinal wholemount cultures provide a system close to the physiological in vivo situation with neuronal interactions and connections still preserved2-5.
In this video article we provide a step by step demonstration of (1) the establishment of in vivo-like organotypic retinal wholemount cultures including dissection peculiarities of embryonic, postnatal and adult murine eyes and (2) a dissociation and cytospin procedure for analysis of neuronal apoptosis and retinal cell proliferation in organotypic wholemounts, e.g. after culture in the presence of exogenously applied recombinant factors.

In vivo-like Organotypic Murine Retinal Wholemount Culture

This video article demonstrates the establishment of organotypic retinal wholemount cultures and a cytospin procedure for analysis of exogenously induced effects. Organotypic retinal wholemount cultures mimic the in vivo situation and significantly facilitate the accessibility of murine retinas for experimental manipulations while circumventing the disadvantages of classical murine animal models.

In vivo-ähnlichen Organotypische Murine Retinal Wholemount Kultur

Targeted ablations of genes and analysis of animal models is the classical strategy for enrolling specific retinal gene function. However, transgenic, ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

Erstellung von Entwicklungs-und Adult Drosophila Brains und retinae für Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

Preparation, Purification, and Characterization of Lanthanide Complexes for Use as Contrast Agents for Magnetic Resonance Imaging

Polyaminopolycarboxylate-based ligands are commonly used to chelate lanthanide ions, and the resulting complexes are 
useful as contrast agents for magnetic resonance imaging (MRI). Many commercially available ligands are especially useful because they contain 
functional groups that allow for fast, high-purity, and high-yielding conjugation to macromolecules and biomolecules via amine-reactive 
activated esters and isothiocyanate groups or thiol-reactive maleimides. While metalation of these ligands is considered common 
knowledge in the field of bioconjugation chemistry, subtle differences in metalation procedures must be taken into account when 
selecting metal starting materials. Furthermore, multiple options for purification and characterization exist, and selection of the most 
effective procedure partially depends on the selection of starting materials. These subtle differences are often neglected in published 
protocols. Here, our goal is to demonstrate common methods for metalation, purification, and characterization of lanthanide complexes 
that can be used as contrast agents for MRI (Figure 1). We expect that this publication will enable biomedical scientists to incorporate 
lanthanide complexation reactions into their repertoire of commonly used reactions by easing the selection of starting materials and 
purification methods.

We demonstrate the metalation, purification, and characterization of lanthanide complexes. The complexes described here can be conjugated to macromolecules to enable tracking of these molecules using magnetic resonance imaging.

Herstellung, Reinigung und Charakterisierung von Lanthanoidkomplexe für die Verwendung als Kontrastmittel für die Magnetresonanz-Tomographie

Polyaminopolycarboxylate-based ligands are commonly used to chelate lanthanide ions, and the resulting complexes are 
useful as contrast agents for ...

The Mouse Cremaster Muscle Preparation for Intravital Imaging of the Microcirculation

Throughout the body, the maintenance of homeostasis requires the constant supply of oxygen and nutrients concomitant with removal of metabolic by-products. This balance is achieved by the movement of blood through the microcirculation, which encompasses the smallest branches of the vascular supply throughout all tissues and organs. Arterioles branch from arteries to form networks that control the distribution and magnitude of oxygenated blood flowing into the multitude of capillaries intimately associated with parenchymal cells. Capillaries provide a large surface area for diffusional exchange between tissue cells and the blood supply. Venules collect capillary effluent and converge as they return deoxygenated blood towards the heart. To observe these processes in real time requires an experimental approach for visualizing and manipulating the living microcirculation.
The cremaster muscle of rats was first used as a model for studying inflammation using histology and electron microscopy post mortem1,2. 
The first in vivo report of the exposed intact rat cremaster muscle investigated microvascular responses to vasoactive drugs using reflected light3. However 
curvature of the muscle and lack of focused illumination limited the usefulness of this preparation. The major breakthrough entailed opening the muscle, 
detaching it from the testicle and spreading it radially as a flat sheet for transillumination under a compound microscope4. While shown to be a valuable 
preparation to study the physiology of the microcirculation in rats5 and hamsters6, the cremaster muscle in mice7 has proven particularly useful in dissecting 
cellular pathways involved in regulating microvascular function8-11 and real-time imaging of intercellular signaling12. 
The cremaster muscle is derived from the internal oblique and transverse abdominus muscles as the testes descend through the inguinal canal13. 
It serves to support (Greek: cremaster = suspender) and maintain temperature of the testes. As described here, the cremaster muscle is prepared as a thin flat sheet 
for outstanding optical resolution. With the mouse maintained at a stable body temperature and plane of anesthesia, surgical preparation involves freeing the muscle 
from surrounding tissue and the testes, spreading it onto transparent pedestal of silastic rubber and securing the edges with insect pins while irrigating it 
continuously with physiological salt solution. The present protocol utilizes transgenic mice expressing GCaMP2 in arteriolar endothelial cells. GCaMP2 is a 
genetically encoded fluorescent calcium indicator molecule12. Widefield imaging and an intensified charge-coupled device camera enable in vivo study of 
calcium signaling in the arteriolar endothelium.

A tissue preparation is described for visualization and experimental manipulation of the living microcirculation. In anesthetized male mice, the thin, highly vascularized cremaster muscle is prepared for intravital microscopy to study microvascular networks including arterioles, capillaries and venules. This preparation is readily adapted for rats and hamsters.

Die Maus Cremaster Muscle Vorbereitung Intravital Imaging der Mikrozirkulation

Throughout the body, the maintenance of homeostasis requires the constant supply of oxygen and nutrients concomitant with removal of metabolic ...

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine parasitoids known to infect the host during the egg stage1. Second, it has a wide range of potential fruit fly hosts. Perhaps due to its life history, F. arisanus has been a successfully used for biological control of fruit flies in multiple tropical regions2-4. One impediment to the wide use of F. arisanus for fruit fly control is that it is difficult to establish a stable laboratory colony5-9. Despite this difficulty, in the 1990s USDA researchers developed a reliable method to maintain laboratory populations of F. arisanus10-12. There is significant interest in F. arisanus biology13,14, especially regarding its ability to colonize a wide variety of Tephritid hosts14-17; interest is especially driven by the alarming spread of Bactrocera fruit fly pests to new continents in the last decade18. Further research on F. arisanus and additional deployments of this species as a biological control agent will benefit from optimizations and improvements of rearing methods. In this protocol and associated video article we describe an optimized method for rearing F. arisanus based on a previously described approach12. The method we describe here allows rearing of F. arisanus in a small scale without the use of fruit, using materials available in tropical regions around the world and with relatively low manual labor requirements.

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus is an egg-larval parasitoid of Tephritid fruit flies that is successfully used in biological control of these important tropical pests. We describe here an optimized protocol for rearing F. arisanus on a small scale using readily available materials.

Ein optimiertes Protokoll für die Aufzucht Fopius arisanus, Fliegt ein Parasitoid von Tephritid Fruit

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

Riesen-Liposomen-Herstellung für Imaging und Patch-Clamp-Elektrophysiologie

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has become the method of choice for introducing DNA into prokaryotes that are not naturally competent. Electroporation is a rapid, efficient, and streamlined transformation method that, in addition to purified DNA and competent bacteria, requires commercially available gene pulse controller and cuvettes. In contrast to the pulsing step, preparation of electrocompetent cells is time consuming and labor intensive involving repeated rounds of centrifugation and washes in decreasing volumes of sterile, cold water, or non-ionic buffers of large volumes of cultures grown to mid-logarithmic phase of growth. Time and effort can be saved by purchasing electrocompetent cells from commercial sources, but the selection is limited to commonly employed E. coli laboratory strains. We are hereby disseminating a rapid and efficient method for preparing electrocompetent E. coli, which has been in use by bacteriology laboratories for some time, can be adapted to V. cholerae and other prokaryotes. While we cannot ascertain whom to credit for developing the original technique, we are hereby making it available to the scientific community.

Rapid Protocol for Preparation of Electrocompetent Escherichia coli and Vibrio cholerae

Electroporation is a commonly employed method for introducing DNA into bacteria in a process known as transformation. Traditional protocols for the preparation of electrocompetent cells are time consuming and labor intensive. This article describes an alternate, rapid, and efficient method for the preparation of electrocompetent cells presently employed by some laboratories.

Rapid-Protokoll für die Herstellung von Elektrokompetente<em&gt; Escherichia coli</em&gt; Und<em&gt; Vibrio cholerae</em

Electroporation has become a widely used method for rapidly and efficiently introducing foreign DNA into a wide range of cells. Electrotransformation has ...

A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle composition. Various time-consuming methods have traditionally been used to study fiber populations in many fields of research. However, recently developed immunohistochemical methods based on myosin heavy chain protein expression provide a quick alternative to identify multiple fiber types in a single section. Here, we present a rapid, reliable and reproducible protocol for improved staining quality, allowing automatic acquisition of whole cross sections and automatic quantification of fiber populations with ImageJ. For this purpose, embedded skeletal muscles are cut in cross sections, stained using myosin heavy chains antibodies with secondary fluorescent antibodies and DAPI for cell nuclei staining. Whole cross sections are then scanned automatically using a slide scanner to obtain high-resolution composite pictures of the entire specimen. Fiber population analyses are subsequently performed to quantify slow, intermediate and fast fibers using an automated macro for ImageJ. We have previously shown that this method can identify fiber populations reliably to a degree of &#177;4%. In addition, this method reduces inter-user variability and time per analyses significantly using the open source platform ImageJ.

Here, we present a protocol for rapid muscle fiber analyses, which allows improved staining quality, and thereby automatic acquisition and quantification of fiber populations using the freely available software ImageJ.

Ein schnelles automatisiertes Protokoll zur Muskelfaser Populationsanalyse in Rattenmuskelquerschnitte Mit Myosin Heavy Chain Immunhistochemie

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle ...