This work was supported by NIH Grant EY 14990.

Part 1: Dissection and Bipolar Cell Isolation
<ol>
<li>Prepare solutions listed in Table 2; The pH of ringers&rsquo; (external) solutions should be adjusted to 7.4 with NaOH and the pH of the internal solution should be adjusted to 7.2 with CsOH. Protect internal solution from light with aluminum foil and keep it at 4 &deg;C until use;</li>
<li>Dark-adapt a goldfish for at least 30 minutes prior to dissection;</li>
<li>While the animal dark-adapts, prepare 5 mL of the hyaluronidase (type V hyaluronidase,1100 units/mL in low Ca2+ ringer&rsquo;s; Sigma, St. Louis, MO) and 10 mL of the L-cysteine (0.5&nbsp;mg/mL in low Ca2+ ringer&rsquo;s) solutions and weigh the papain (lyophilized powder, 40&nbsp;units/mL; Sigma, St. Louis, MO) for 5 mL of the digestion solution;</li>
<li>Euthanize the goldfish by quick decapitation with surgical scissors and destroy the brain and spinal cord with a #11 scalpel blade;</li>
<li>Remove the eyes by destroying the extra-ocular muscles with the help of a #7 curved Dumont forceps and cutting the optic nerve with iris scissors;</li>
<li>Place one eye bulb on a piece of filter paper and puncture the scleral limbus with the tip of a #11 scalpel blade;</li>
<li>Introduce the blade of a pair of vannas scissors inside the punctured whole and cut the whole anterior segment away;</li>
<li>Place a small piece of filter paper on top of the remaining optic cup and exert some pressure in order to have the paper soak with vitreous humor;</li>
<li>Lift the filter paper with the retina attached to it and cut the optic nerve with a par of vannas scissors;</li>
<li>Place the filter paper containing the retina in a 35 mm plastic culture dish with hyaluronidase solution and peel off the retina from the filter paper with the help of #7 Dumont tweezers;</li>
<li>Cut the retina into 4-6 pieces with half of an industrial carbon steel single-edged blade and let it sit in the hyaluronidase solution for 20 minutes;</li>
<li>While waiting for hyaluronidase to take effect, add 5 mL of the L-cysteine solution to the papain and let it sit until the liquid becomes transparent (approximately 5-10 minutes);</li>
<li>Wash the pieces of retina 3x in low Ca2+ Ringer&rsquo;s and let them sit in the papain solution for 30-35 minutes;</li>
<li>Wash the pieces of retina 3x in low Ca2+ Ringer&rsquo;s and store them until use at 4 &deg;C in a 35&nbsp;mm plastic culture dish containing low Ca2+ Ringer&rsquo;s;</li>
<li>To dissociate the cells, put a piece of retina in a microcentrifuge tube containing 500 mL of low Ca2+ Ringer&rsquo;s and slowly triturate the retina by pipetting it up and down with a glass dissociation pipette, carefully not to produce any air bubbles. Dissociation pipettes are fabricated by heating up the tip of a glass Pasteur pipette with a Bunsen burner and slightly bending it with the help of anatomical forceps;</li>
<li>Plate the isolated cells by adding a drop of the retinal suspension to a home-made recording chamber previously filled with 2 mL of low Ca2+ ringer&rsquo;s. The chamber consists of the bottom half of a 35-mm plastic culture dish with a circular whole in the middle and a circular coverslip of 1.78 refractive index glass (PlanOptik, Germany) glued to the bottom with a silicon elastomer (Sylgard 184; Dow Corning, Midland, MI).</li>
</ol>
Part 2: Bipolar Cell Loading and Wash Out
TIRF imaging of synaptic vesicles is best carried out using an objective-type TIRFM microscope with a very high NA objective and a sensitive camera. For our experiments, we choose to use a 1.65 NA objective (Apo x100 O HR, N.A. 1.65, Olympus, Japan) with an EMCCD (Cascade 512B, Roper Scientific, Tucson, AZ). The use of the very high NA objective necessitates the use of high refractive glass coverslips and immersion fluid (di-iodomethane with sulfur). Under our conditions, excitation light is limited to an exponentially decaying field with a length constant of approximately 50 nm.
<ol>
<li>Add a drop of high refractive index liquid (M series, refractive index = 1.7800, Cargille Labs, Cedar Grove, NJ) to the microscope objective;</li>
<li>Place the recording chamber carefully on top of the microscope objective and carefully mount ground electrode and superfusion exit pipe to chamber;</li>
<li>Let chamber sit on microscope for 10-20 minutes to allow cells to sink and adhere to the bottom;</li>
<li>In the meantime, prepare 5 mL of a 1mM trolox&reg; ((&plusmn;)-6-Hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid; Sigma, St. Louis, MO) solution in high K+ ringer&rsquo;s. Sonicate until dissolved;</li>
<li>Prepare 15 mL of ADVASEP-7 washing solution: 1 mM ADVASEP-7 (Sigma, St. Louis, MO) in low Ca2+ ringer&rsquo;s. Note that ADVASEP-7 use is optional and can be omitted if desired;</li>
<li>Purge the superfusion lines and add ADVASEP-7 washing solution, low Ca2+ ringer&rsquo;s and control ringer&rsquo;s to the superfusion system; </li>
<li>Pull loading pipettes from thin-walled borosilicate glass (Kwik-Fil&reg; TW150-3; WPI, Sarasota, FL). Puffer pipette resistances are in the 1.5-2.5 M&Omega; range;</li>
<li>Prepare the FM1-43&reg; (N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium dibromide, &ldquo;special packaging&rdquo;; Invitrogen, Carlsbad, CA) solutions. First, make a 1 mM stock by adding 160 &mu;L distilled water to one vial (100 mg) of FM1-43&reg;. This stock can be kept at 4 &deg;C for up to one week. Then, add 5 &mu;L of FM1-43&reg; to 1 mL high K+ ringer&rsquo;s + 1mM trolox&reg;. Protect solution from light with aluminum foil and keep it at 4 &deg;C until use; </li>
<li>Turn the microscope bright field light on and search for intact bipolar cells. Slightly tap the microscope to make sure that the neurons are firmly attached to the bottom of the chamber;</li>
<li>Position the superfusion pen close to the cell of interest and continuously perfuse the preparation with low Ca2+ ringer&rsquo;s;</li>
<li>Turn off the room lights and add a red long pass filter (i.e. RG630; Schott, Germany) to the optical path to minimize excitation of the FM dye;</li>
<li>Fill a loading pipette with 10 &mu;L of the FM dye solution, mount the pipette in the micromanipulator and lower the pipette onto the preparation without overpressure until it is at the same focal plane as the bipolar cell you want to load. Make sure you have at least two electrode holders: the one used for the FM dye cannot be used for patch clamping, or else it may contaminate the intracellular solution;</li>
<li>Position the puffer opening at a distance of around 10 &mu;m from the axon terminal, turn the superfusion system off and puff the dye solution for 10 seconds by turning the pipette overpressure on;</li>
<li>Turn the overpressure off and, without moving the pipette, wait for 30 seconds;</li>
<li>Turn the superfusion on and bathe the chamber in ADVASEP-7 solution for 5 minutes. In the meantime, remove the puffer pipette from the bath;</li>
<li>After 5 minutes, switch to low Ca2+ ringer&rsquo;s and perfuse the chamber for 25-30 minutes to allow removal of excess dye.</li>
</ol>
Part 3: Patch Clamping and TIRFM Imaging
<ol>
<li>While the preparation is washing out, pull patch pipettes from thick-walled borosilicate glass (B150-86-10; Sutter Instrument Company, Novato, CA). Patch pipette resistances are in the 8-10 M&Omega; range;</li>
<li>After wash out is complete, place the axon terminal in the center of the field of view;</li>
<li>Fill a patch pipette with 7&mu;L of internal solution, tap the pipette to get rid of air bubbles, coat the pipette with molten dental wax (Sticky Wax; Kerr Corporation, Orange, CA) and mount it in the micromanipulator;</li>
<li>Turn the pipette overpressure on and lower the pipette slowly onto the preparation. Check pipette resistance in the amplifier and correct pipette offsets and capacitance with the respective amplifier controls;</li>
<li>Switch the superfusion to control ringer&rsquo;s. To create a gigaseal between the pipette and the cell, slightly touch the tip of the electrode against the cell body and turn the pipette overpressure off while gently applying negative pressure to the electrode;</li>
<li>While sealed, choose &ldquo;whole cell&rdquo; mode in the amplifier and set the cell holding potential to -60 mV;</li>
<li>Choose a region of interest with the imaging software that encompasses the whole axon terminal, turn the bright field light off and, by briefly exposing (30 ms) the terminal to the 488 nm laser, find the right focal plane for TIRF imaging;</li>
<li>Break into the cell by using the &ldquo;zap&rdquo; command of the amplifier while applying slightly negative pressure to the pipette;</li>
<li>Correct for cell capacitance and series resistance, and then apply the voltage protocol of interest while imaging the movement of synaptic vesicles. It is at this stage important to have the voltage protocol synchronized to the camera frame rate. In our case, each frame is 30 ms long, so the voltage changes occur in multiples of this value (i.e., every 300 ms or 10 frames);</li>
<li>Wait at least 40 seconds between trials to allow for recovery;</li>
<li>To check the position of synaptic ribbons, take pictures while turning the 561 nm laser on.</li>
</ol>
<img src="/files/ftp_upload/1305/fig1.jpg" alt="figure 1" /> Figure 1: The experimental setup. A 488 nm laser (blue) is focused to the periphery of the back focal plane of the objective and suffers total internal reflection when it reaches the glass-aqueous medium interface. The electromagnetic field generated by the reflected beam excites the fluorophore loaded into the synaptic vesicles closest to the bottom of the glass chamber, which then fluoresce (green). The fluorescent light is then guided to the observer&rsquo;s eye (depicted) or to a CCD camera. The membrane potential of the imaged cells is controlled simultaneously by patch-clamping them. This approach allows the study of the relationship between incoming signals (the membrane voltage) and the neuronal output (exocytosis).
<img src="/files/ftp_upload/1305/fig2.jpg" alt="figure 2" /> Figure 2: Typical results. Left: bright field image of an isolated goldfish bipolar cell. Upper right: TIRF image of the synaptic vesicles in the bipolar cell axon terminal loaded with FM 1-43&reg; and imaged with the 488 nm laser (FM dye). Bottom right: image of the same terminal after patching the cell and imaging the axon terminal with the 561 nm laser. Synaptic ribbons are labeled by the rhodamine-based RIBEYE-binding peptide (Rpep_rhod).
Table 1: Specific Reagents and Equipment.
<table border="1" cellspacing="0" cellpadding="0">
<tbody>
<tr>
<td> Name</td>
<td>Type</td>
<td>Manufacturer</td>
<td>Catalog</td>
<td>Comment</td>
</tr>
<tr>
<td>-</td>
<td>Air Table</td>
<td>Newport Corporation</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>IX70</td>
<td>Inverted Microscope</td>
<td>Olympus</td>
<td>-</td>
<td>Equipped with a tungsten lamp for bright field and a lateral opening port for TIRF</td>
</tr>
<tr>
<td>TH4-100</td>
<td>Lamp Power Source</td>
<td>Olympus</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>FF498_581-Di01</td>
<td>Dichroic Filter</td>
<td>Semrock</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>NF01-405_488_568</td>
<td>Emission Filter</td>
<td>Semrock</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Apo x100 O HR</td>
<td>Objective</td>
<td>Olympus</td>
<td>-</td>
<td>N.A. 1.65</td>
</tr>
<tr>
<td>RG630</td>
<td>Red Glass Filter</td>
<td>Schott</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>-</td>
<td>488&nbsp;nm Laser</td>
<td>Coherent</td>
<td>-</td>
<td>Use at minimum power</td>
</tr>
<tr>
<td>-</td>
<td>Shutter</td>
<td>Uniblitz</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>VMM-D1</td>
<td>Shutter Driver</td>
<td>Uniblitz</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>-</td>
<td>561&nbsp;nm Laser</td>
<td>Melles Griot</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>-</td>
<td>Shutter</td>
<td>Uniblitz</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>VMM-D3</td>
<td>Shutter Driver</td>
<td>Uniblitz</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Perfusion Pressure Kit</td>
<td>Superfusion System</td>
<td>Automate Scientific</td>
<td>09-04</td>
<td>-</td>
</tr>
<tr>
<td>Perfusion Pen</td>
<td>Superfusion System</td>
<td>Automate Scientific</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Valvelink 8</td>
<td>Superfusion System Controller</td>
<td>Automate Scientific</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Cascade 512B</td>
<td>EM CCD Camera</td>
<td>Roper Scientific</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Metamorph 7.1</td>
<td>Imaging Software</td>
<td>Molecular Devices</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>EPC-9</td>
<td>Patch Clamp Amplifier</td>
<td>HEKA Elektronik</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Pulse</td>
<td>Amplifier Software</td>
<td>HEKA Elektronik</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>MP-285</td>
<td>Micromanipulator</td>
<td>Sutter Instrument</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>-</td>
<td>Electrode Holder</td>
<td>HEKA Elektronik</td>
<td>-</td>
<td>For 1.5 mm O.D. glass, 2 units</td>
</tr>
<tr>
<td>Kwik-Fil&reg; TW150-3</td>
<td>Borosilicate Capillary Glass</td>
<td>WPI</td>
<td>-</td>
<td>Without filament</td>
</tr>
<tr>
<td>B150-86-10</td>
<td>Borosilicate Capillary Glass</td>
<td>Sutter Instrument</td>
<td>-</td>
<td>With filament</td>
</tr>
<tr>
<td>P-97</td>
<td>Microelectrode Puller</td>
<td>Sutter Instrument</td>
<td>-</td>
<td>Equipped with 3x3 box filament and environmental chamber</td>
</tr>
<tr>
<td>-</td>
<td>Pressure Vacuum Air Pump</td>
<td>Thomas Scientific</td>
<td>7893B05</td>
<td>Creates vacuum to remove liquids from chamber and overpressure for pipettes</td>
</tr>
<tr>
<td>MatLab R2008a</td>
<td>Analysis Software</td>
<td>MathWorks</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>353001</td>
<td>35&nbsp;mm Plastic Culture Dishes</td>
<td>Falcon</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>-</td>
<td>High Refractive Index Glass</td>
<td>PlanOptik</td>
<td>-</td>
<td>Refractive index488 nm = 1.78</td>
</tr>
<tr>
<td>Series M</td>
<td>High Refractive Index Liquid</td>
<td>Cargille Labs</td>
<td>-</td>
<td>Refractive index = 1.78</td>
</tr>
<tr>
<td>Sylgard 184</td>
<td>Silicon Elastomer Kit</td>
<td>Dow Corning</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Glutathione</td>
<td>Tripeptide</td>
<td>EMD Chemicals</td>
<td>&nbsp;</td>
<td>Free radical scavenger</td>
</tr>
<tr>
<td>Hyaluronidase</td>
<td>Enzyme</td>
<td>Sigma</td>
<td>H6254</td>
<td>Type V</td>
</tr>
<tr>
<td>L-cysteine</td>
<td>Amino Acid</td>
<td>Fluka</td>
<td>30090</td>
<td>Activates papain</td>
</tr>
<tr>
<td>Papain</td>
<td>Enzyme</td>
<td>Fluka</td>
<td>76220</td>
<td>From Carica papaya</td>
</tr>
<tr>
<td>Trolox&reg;</td>
<td>Soluble Vitamin E</td>
<td>Sigma</td>
<td>56510</td>
<td>Free radical scavenger</td>
</tr>
<tr>
<td>ADVASEP-7</td>
<td>Sulfonated B-Cyclodextrin Derivative</td>
<td>Sigma</td>
<td>A3723</td>
<td>Reduces FM 1-43&reg; background fluorescence</td>
</tr>
<tr>
<td>FM 1-43&reg;</td>
<td>Fluorescent Dye</td>
<td>Invitrogen</td>
<td>T35356</td>
<td>&ldquo;Special packaging&rdquo;</td>
</tr>
<tr>
<td>Sticky Wax</td>
<td>Pipette Coating Agent</td>
<td>Kerr Corporation</td>
<td>-</td>
<td>Decreases pipette capacitance</td>
</tr>
</tbody>
</table>
Table 2: Physiological Solutions Used in This Study.
<table border="1" cellspacing="0" cellpadding="0">
<tbody>
<tr>
<td> Substance</td>
<td>Low Ca2+ Ringer&rsquo;s</td>
<td>Control Ringer&rsquo;s</td>
<td>High K+ Ringer&rsquo;s</td>
<td>Internal Solution</td>
</tr>
<tr>
<td>NaCl</td>
<td>120 mM</td>
<td>120 mM</td>
<td>97.5 mM</td>
<td>-</td>
</tr>
<tr>
<td>KCl</td>
<td>2.5 mM</td>
<td>2.5 mM</td>
<td>25 mM</td>
<td>-</td>
</tr>
<tr>
<td>MgCl2</td>
<td>1 mM</td>
<td>1 mM</td>
<td>1 mM</td>
<td>4 mM</td>
</tr>
<tr>
<td>CaCl2</td>
<td>0.5 mM</td>
<td>2.5 mM</td>
<td>2.5 mM</td>
<td>-</td>
</tr>
<tr>
<td>HEPES</td>
<td>10 mM</td>
<td>10 mM</td>
<td>10 mM</td>
<td>10 mM</td>
</tr>
<tr>
<td>EGTA</td>
<td>0.75 mM</td>
<td>-</td>
<td>-</td>
<td>0.5 mM</td>
</tr>
<tr>
<td>Glucose</td>
<td>10 mM</td>
<td>10 mM</td>
<td>-</td>
<td>-</td>
</tr>
<tr>
<td>Glutathione</td>
<td>2 mM</td>
<td>2 mM</td>
<td>-</td>
<td>1 mM</td>
</tr>
<tr>
<td>CH3CsO3S*</td>
<td>-</td>
<td>-</td>
<td>-</td>
<td>100 mM</td>
</tr>
<tr>
<td>TEACl</td>
<td>-</td>
<td>-</td>
<td>-</td>
<td>10 mM</td>
</tr>
<tr>
<td>ATP-Mg</td>
<td>-</td>
<td>-</td>
<td>-</td>
<td>10 mM</td>
</tr>
<tr>
<td>GTP-Li</td>
<td>-</td>
<td>-</td>
<td>-</td>
<td>1 mM</td>
</tr>
<tr>
<td>Rpep-rhod**</td>
<td>-</td>
<td>-</td>
<td>-</td>
<td>5 mM</td>
</tr>
<tr>
<td>Volume</td>
<td>200 mL</td>
<td>100 mL</td>
<td>5 &mu;L</td>
<td>100 &mu;L</td>
</tr>
</tbody>
</table>
* Cesium methanesulfonate. ** RIBEYE-binding peptide: rhodamine+EQTVPVDLSVARDR-cooh (mw 1997.75).

<ol>
	<li>Axelrod, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Total+internal+reflection+fluorescence+microscopy+in+cell+biology.">Total internal reflection fluorescence microscopy in cell biology.</a> Traffic. 2, 764-774 (2001).</li><li>Zenisek, D., Steyer, J. A., Almers, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transport,+capture+and+exocytosis+of+single+synaptic+vesicles+at+active+zones.">Transport, capture and exocytosis of single synaptic vesicles at active zones.</a> Nature. 406, 849-854 (2000).</li><li>Zenisek, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vesicle+association+and+exocytosis+at+ribbon+and+extraribbon+sites+in+retinal+bipolar+cell+presynaptic+terminals.">Vesicle association and exocytosis at ribbon and extraribbon sites in retinal bipolar cell presynaptic terminals.</a> Proc. Natl. Acad. Sci. U. S. A. 105, 4922-4927 (2008).</li><li>Zenisek, D., Horst, N. K., Merrifield, C., Sterling, P., Matthews, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+synaptic+ribbons+in+the+living+cell.">Visualizing synaptic ribbons in the living cell.</a> J. Neurosci. 24, 9752-9759 (2004).</li><li>Axelrod, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+imaging+of+surface+fluorescence+with+very+high+aperture+microscope+objectives.">Selective imaging of surface fluorescence with very high aperture microscope objectives.</a> J. Biomed. Opt. 6, 6-13 (2001).</li><li>Rouze, N. C., Schwartz, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Continous+and+transient+vesicle+cycling+at+a+ribbon+synapse.">Continous and transient vesicle cycling at a ribbon synapse.</a> J. Neurosci. 18, 8614-8624 (1998).</li></ol>

The advantages of objective-type TIRF microscopy are that 1) it provides excellent optical sectioning by restricting excitation light to a narrow region within the focal plane of the objective, thereby minimizing out-of-focus light; 2) since light drops exponentially with distance, movement in a vertical direction can be monitored as a change in fluorescence intensity; 3) efficient light collection through the high numerical aperture objective1,5.
The main drawback of the technique...

imaging exocytosis in retinal bipolar cells with tirf microscopy

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective imaging of fluorescent molecules that are closest to a high refractive index substance such as glass1. In this article, we apply this technique to image exocytosis of synaptic vesicles in retinal bipolar cells isolated from the goldfish retina. These neurons are very suitable for this kind of study due to their large axon terminals. By simultaneously patch clamping the bipolar cells, it is possible to investigate the relationship between pre-synaptic voltage and synaptic release2,3. Synaptic vesicles inside the bipolar cell terminals are loaded with a fluorescent dye (FM 1-43&reg;) by co-puffing the dye and a ringer solution containing a high K+ concentration onto the synaptic terminals. This depolarizes the cells and stimulates endocytosis and consequent dye uptake into the glutamatergic vesicles. After washing the excess dye away for around 30 minutes, cells are ready for being patch clamped and imaged simultaneously with a 488 nm laser. The patch pipette solution contains a rhodamine-based peptide that binds selectively to the synaptic ribbon protein RIBEYE4, thereby labeling ribbons specifically when terminals are imaged with a 561 nm laser. This allows the precise localization of active zones and the separation of synaptic from extra-synaptic events.

Watch this Scientific Journal Video about Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy at JoVE.com

Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

In this video, we demonstrate how to label and visualize single synaptic vesicle exocytosis and trafficking in goldfish retinal bipolar cells using total internal reflectance fluorescence (TIRF) microscopy.

Total internal reflectance fluorescence (TIRF) microscopy is a technique that allows the study of events happening at the cell membrane, by selective ...

imaging-exocytosis-in-retinal-bipolar-cells-with-tirf-microscopy

Research

JoVE Journal

Biology

11.6K vistas.  Yale University School of Medicine. En este video, que muestra cómo la etiqueta y visualizar solo exocitosis de vesículas sinápticas y el tráfico en las células de la retina con peces de colores bipolar total de fluorescencia de reflexión interna (TIRF) microscopía.

Video: Imaging Exocytosis in Retinal Bipolar Cells with TIRF Microscopy

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...

Imaging FITC-dextran as a Reporter for Regulated Exocytosis

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated exocytosis is fundamental for intercellular communication and is a key mechanism for the secretion of neurotransmitters, hormones, inflammatory mediators, and other compounds, by a variety of cells. At least three distinct mechanisms are known for regulated exocytosis: full exocytosis, where a single SG fully fuses with the plasma membrane, kiss-and-run exocytosis, where a single SG transiently fuses with the plasma membrane, and compound exocytosis, where several SGs fuse with each other, prior to or after SG fusion with the plasma membrane. The type of regulated exocytosis undertaken by a cell is often dictated by the type of secretory trigger. However, in many cells, a single secretory trigger can activate multiple modes of regulated exocytosis simultaneously. Despite their abundance and importance across cell types and species, the mechanisms that determine the different modes of secretion are largely unresolved. One of the main challenges in investigating the different modes of regulated exocytosis, is the difficulty in distinguishing between them as well as exploring them separately. Here we describe the use of fluorescein isothiocyanate (FITC)-dextran as an exocytosis reporter, and live cell imaging, to differentiate between the different pathways of regulated exocytosis, focusing on compound exocytosis, based on the robustness and duration of the exocytic events.

Here we detail a method for live cell imaging of regulated exocytosis. This method utilizes FITC-dextran, which accumulates in lysosome-related organelles, as a reporter. This simple method also allows distinguishing between different modes of regulated exocytosis in cells that are difficult to manipulate genetically.

Regulated exocytosis is a process by which cargo, which is stored in secretory granules (SGs), is released in response to a secretory trigger. Regulated ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...