We thank J. Chen for providing the caged fluorescein-dextran synthesis protocol. This research was supported by the March of Dimes award 5-FY09-78, the American Heart Association award 09BGIA2090075, and the NIH / NHLBI award R01 HL107369-01 to S.S. A special thank you to C. Closson for his microscopy assistance.

1. Synthesis of caged fluorescein-dextran (adapted from Ref. 14) 
<ol>
<li>Prepare a 0.1 M solution of sodium borate diluted in ddH2O.</li>
<li>Measure 4 mg of 10 kDa aminodextran and add it to the CMNB-caged fluorescein tube.</li>
<li>Add 500 &mu;L of the prepared sodium borate solution (step 1.1) to the CMNB-caged fluorescein tube. Cap the tube and vortex for 30 seconds to dissolve the mixture.</li>
<li>Allow the mixture to react overnight on a nutator at room temperature. Cover the tube with metal foil to prevent exposure of the mixture to ambient light.</li>
<li>Obtain a Zeba desalt spin column and twist off the bottom cap. Mark a dot on the column, which will be used to orient the column when centrifuging. Place the column in a 15 mL conical Falcon tube. Note: The solution in the spin column is column buffer.</li>
<li>Loosen the top cap on the desalt spin column. Place the 15 mL conical Falcon tube that houses the desalt spin column in a centrifuge. Orient the spin column with the marked dot (step 1.4) facing the center of the centrifuge rotor. Centrifuge the tube at 1000 x g for 2 minutes at room temperature.</li>
<li>After centrifugation, remove the desalt spin column from the Falcon tube. Discard both the collected flow through and the 15 mL conical tube. Obtain a new 15 mL conical Falcon tube, and place the desalt spin column in the new Falcon tube. Note: After centrifuging, the column resin bed should appear compacted. Use the column immediately.</li>
<li>Obtain the reacted mixture (caged fluorescein-dextran) in step 1.3. Remove the cap from the spin column and aspirate the reacted mixture and apply it slowly to the center of the desalt spin column. After applying the reacted mixture, replace the cap on the spin column.</li>
<li>Place the Falcon tube that houses the desalt spin column in a centrifuge. As done in step 1.5, orient the spin column with the marked dot facing the center of the centrifuge rotor. Centrifuge the tube at 1000 x g for 2 minutes at room temperature.</li>
<li>After centrifugation, a yellow mixture (caged fluorescein-dextran) will be collected (approximately 400 &mu;L) in the 15 mL Falcon tube. Transfer the caged fluorescein-dextran mixture to a 1.5 mL microcentrifuge tube.</li>
<li>Immediately cover the tube with metal foil to prevent exposure of the mixture to ambient light. Lyophilize the caged fluorescein-dextran mixture in a speedvac. Lyophilization of 350 &mu;L should take approximately 4 hours.</li>
<li>After complete lyophilization, resuspend the powder (about 3 mg) in ddH2O to a final concentration of approximately 1 % w/v. Mix by briefly vortexing to suspend the powder.</li>
<li>Aliquot the caged fluorescein-dextran into separate metal foil covered tubes, and store them at 
-20 &deg;C.</li>
</ol>
2. Harvesting of embryos and microinjection of caged fluorescein-dextran
* This procedure uses the zebrafish as the animal model system. 
<ol>
<li>On the day before injections, set up zebrafish crosses either 1:1 male-to-female or 2:1 male-to-female in breeding tanks with dividers.</li>
<li>The next morning change the breeding tank water and remove the dividers. Immediately prepare a 5 &mu;L 1/10 diluted working solution of caged fluorescein-dextran in either 1 X Danieau media (see Preparation of Solutions) or ddH2O. Cover the working solution tube with metal foil to prevent exposure to light.</li>
<li>Pipette the prepared caged fluorescein-dextran solution (step 2.2) into the needle. Before cutting the needle tip to an appropriate size under the dissection microscope, place a piece of transparent yellow filter in front of the white light source. The yellow filter will prevent spontaneous uncaging of the fluorescein-dextran during injections. When looking through the ocular eye piece, the light source should appear yellow.</li>
<li>Collect embryos and pipette them into a microinjection mold tray. Set the microinjection time to deliver 3 to 4 nL of caged fluorescein-dextran. Under the dissection microscope, orient the embryos with forceps and inject (3 to 4 nL) caged fluorescein-dextran into the base of the blastomere cells (blastomere-yolk interface). The embryo stage for injection should be 1- to 2-cell stage.</li>
<li>After injection, remove the embryos from the microinjection mold tray and place them in a clean petri dish containing fresh fish water. Methylene blue may be added to the fish water to prevent fungal growth at a final concentration of at 0.02 % w/v. Cover the petri dish with metal foil to prevent uncaging of the injected caged fluorescein-dextran.</li>
<li>Raise the injected embryos to a stage when the cells of interest need to be photoactivated. For photoactivation, prepare a 0.6 % low melting agarose solution in ddH2O. Depending on the embryo stage, Tricaine (MS222) can be added to the low melting agarose. To do this, dilute 0.02% Tricaine in low melting agarose to a final concentration of 0.0014 %.</li>
<li>As soon as the embryos reach the appropriate developmental stage, dechorionate the embryos in 1 X Danieau media with forceps. When dechorionating, make sure that a transparent yellow filter is placed in the path of the white light source (see step 2.3).</li>
<li>Heat the 0.6 % low melting agarose solution to 37 &deg;C. Pipette 1 mL of low melting agarose into the well of a LabTek chambered coverglass slide. Carefully mount 3 to 4 embryos in low melting agarose and orient them with forceps with the cells to be photoactivated facing downward against the bottom coverslip. </li>
<li>Allow the agarose to solidify. After solidification, pipette 1 mL of 1 X Danieau media over the agarose.</li>
<li>Repeat steps 2.8 to 2.9 for more embryos.</li>
</ol>
3. Two-photon activation of caged fluorescein-dextran
* This procedure assumes that the embryo expresses reporter GFP. The embryo is viewed using the argon ion (488 nm) laser source attached to the LSM 510 META NLO microscope. A single GFP positive cell is selected and photoactivated using the Mai Tai laser (femtosecond laser) coupled to the LSM 510. The procedure is the same for non-fluorescent embryos. Alternatively, white light can be used to find the cell to be activated, followed by activation using the Mai Tai laser source.
<ol>
<li>Load the LSM 510 software. Place a transparent yellow filter in the white light beam path.</li>
<li>Select Scan New Image and click Start Expert Mode.</li>
<li>Select the Laser tab. Turn on the argon ion (488 nm) and Mai Tai (femtosecond) laser sources.</li>
<li>Set the Mai Tai wavelength to 740 nm.</li>
<li>Select the config tab. Set the optical path configuration for the argon ion and Mai Tai laser.</li>
<li>Select the Scan tab. Change the objective to 20X/0.8.</li>
<li>Set the max scan speed by clicking Max.</li>
<li>Set Mode, Method, and Number to Line, Mean, and 1.</li>
<li>Under the Channels tab deselect all laser sources except for the Mai Tai.</li>
<li>Place a power meter in the optical beam path.</li>
<li>Select the Cont button to activate continuous scanning.</li>
<li>Adjust the transmission % until the power meter reads an average laser power of 47 mW.</li>
<li>Stop the scan by selecting the Stop button.</li>
<li>Under the Channels tab deselect the Mai Tai laser source and select the argon ion (488 nm).</li>
<li>Select the Fast xy button. Under the Channels tab adjust the pinhole, detector gain, amplifier offset, and amplifier gain for the argon ion laser to achieve the best image quality.</li>
<li>Using the stage controller, find and focus on the cell to activate.</li>
<li>Click the Stop button.</li>
<li>Using the zoom tool, zoom in on the activated cell until the zoom factor under the Mode tab displays 50 to 70. </li>
<li>Stop the scan by selecting the Stop button. </li>
<li>Under the Channels tab deselect the argon ion laser (488 nm) and select the Mai Tai laser.</li>
<li>Under the Mode tab set the scan speed to 31.46 sec.</li>
<li>Select the Single button to start the scan for two-photon activation of caged fluorescein-dextran of the selected cell. </li>
<li>After activation, deselect the Mai Tai laser under the Channels tab, and reselect the argon ion laser (488 nm). </li>
<li>Set the zoom factor under Mode to 1, and click the Max button under the Speed heading to set the fastest scan speed.</li>
<li>Select Fast xy to review the photoactivated region. Check that the photoactivated region is not laser ablated. For more information about laser ablation, please refer to the discussion section.</li>
<li>Repeat the above steps for other embryos.</li>
</ol>
4. Immunodetection of uncaged fluorescein-dextran (adapted from Ref. 16)
<ol>
<li>After photoativation, place a transparent yellow filter in front of the white light source and manually remove each embryo from low melting agarose using sharp forceps. Place all removed embryos in a new petri dish containing fresh 1 X Danieau media. Cover the petri dish with metal foil to prevent exposure of embryos to light. </li>
<li>If needed, rear the embryos to the desired developmental stage. In the meantime, prepare a 4 % paraformaldehyde solution (see Preparation of Solutions). Aliquot 1.5 mL of prepared paraformaldehyde into a microcentrifuge tube.</li>
<li>After the embryos have reached the developmental stage of interest, pipette the embryos into the microcentrifuge tube (from step 4.2). Cover the tube with metal foil to prevent exposure to light.</li>
<li>Nutate the tube overnight at 4&deg;C. For the next day prepare graded ethanol (EtOH) solutions in phosphate buffer saline (PBS) and ddH2O; 30 % EtOH:PBS, 50 % EtOH:PBS, 70 % EtOH:ddH2O, and 100 % EtOH.</li>
<li>The next day, remove the embryos from 4&deg;C. Remove the metal foil covering the tube and quickly aspirate off paraformaldehye (leave behind a small volume that is enough to cover the embryos). Pipette 1.5 mL of 30 % EtOH:PBS into the tube. Re-cover the tube with metal foil. Nutate the tube at room temperature for 10 mins.</li>
<li>Repeat step 4.5 using 50, 70, and 100 % graded EtOH solutions. After the 100 % EtOH wash, rinse the embryos again in 100 % EtOH for 10 mins.</li>
<li>After the washes, store the embryos overnight at -20 &deg;C. For the next day prepare phosphate buffered tween (PBT; see Preparation of Solutions), 5 X blocking buffer (see manufacturer protocol for preparation), maleic acid (protocol for preparation can be found in the blocking buffer protocol), 10 mg/ml proteinase K, and a 10 X antibody solution (Anti-fluorescein-AP; see Preparation of Solutions).</li>
<li>Also prepare 2 % lamb serum (LS; see Preparation of Solutions) diluted in PBT. Store the antibody at 4 &deg;C overnight shaking on a nutator. The 2 % LS can be kept at 4 &deg;C.</li>
<li>The next day remove the embryos from -20 &deg;C. Remove the metal foil covering the tube and quickly aspirate off 100 % EtOH. Add 1.5 mL of 70 % EtOH:ddH2O to the tube. Re-cover the tube with metal foil. Nutate the tube at room temperature for 10 mins.</li>
<li>Repeat step 4.8 using 50 and 30 % graded EtOH solutions, and 100 % PBT. After the 100 % PBT wash, rinse the embryos 3 more times in 100 % PBT for 10 mins. </li>
<li>If the embryos are older than 24 hours, proteinase K treat them. If not, proceed to step 4.13. To do this, dilute the prepared proteinase K (step 4.7) 1:1000 in PBT. Incubate the embryos in proteinase K for 10 mins, or longer if the embryos are older. Keep the embryos covered with metal foil to prevent exposure to light.</li>
<li>After proteinase K treatment, wash the embryos 5 times in PBT for 10 mins. After washing, fix the embryos in 4 % paraformaldehyde for 30 mins at room temperature on a nutator. Afterward, immediately wash the embryos 5 times in PBT for 10 mins. Keep the embryos covered with metal foil to prevent exposure to light.</li>
<li>Make blocking buffer by diluting 5 X blocking buffer in maleic acid buffer to 1 X. Remove the embryos from PBT and place them in 1 X blocking buffer. Cover the embryos with metal foil and nutate them at room temperature for 5 to 6 hours.</li>
<li> After 5 to 6 hours of blocking, heat shock the embryos at 65 &deg;C for 15 to 20 mins.</li>
<li>Obtain the antibody solution and the 2 % LS prepared the previous day. Centrifuge the antibody solution at max rpm. Transfer the aqueous phase to the 2 % LS tube. Invert the tube to mix.</li>
<li>Transfer the embryos from blocking buffer to the LS-antibody mixture. Keep the embryos covered with metal foil to prevent exposure to light. Nutate the embryos at 4 &deg;C overnight.</li>
<li>The next day wash the embryos at room temperature 6 times for 15 mins in PBT. Prepare AP buffer (see Preparation of Solutions).</li>
<li> Wash the embryos at room temperature 2 times for 10 mins in AP buffer. Prepare NBT/BCIP developing solution (see Preparation of Solutions).</li>
<li>Transfer the embryos to NBT/BCIP. Allow the stain to develop in the dark.</li>
<li>Stop the development by washing the embryos 3 times in PBT for 5 mins each.</li>
<li>For image acquisition mount the embryos in 0.6 % low melting agarose or 3 % methylcellulose, and acquire images using an inverted or upright compound microscope.</li>
<li>Photoactivated samples can be stored for future analysis (staining remains stable) by dehydrating them in a graded EtOH wash. Follow steps 4.4 through 4.7 for dehydrating the samples.</li>
</ol>
5. Representative Results:
<img src="/files/ftp_upload/3172/3172fig1.jpg" alt="Figure 1"/> Figure 1. Photoactivation of caged fluorescein-dextran. (A,B) Brightfield and fluorescence image of a lateral view zebrafish embryo at the 13/14-somite stage before photoactivation. (C,D) The embryo was photoactivated at the13/14-somite stage in one of the somites (arrow) with a wavelength of 740 nm, scan time of 31 secs, and an average laser power of 47 mW. Post-activation, (d) fluorescence from uncaged fluorescein-dextran is observed. Orientation: Dorsal (right), Ventral (left). Scale bar 100 &mu;m.
<img src="/files/ftp_upload/3172/3172fig2.jpg" alt="Figure 2"/> Figure 2. Immunodetection of caged fluorescein-dextran. Figure 2 depicts the immunostaining of uncaged fluorescein-dextran in an 18/19 hpf zebrafish embryo. At the 10-somite stage a small region of the embryo was activated in the lateral plate mesoderm using the same laser parameters stated in Figure 1. Using the immunodetection procedure, the arrow identifies the activated region with minimal background staining. Orientation: Dorsal (top), Ventral (bottom). Scale bar 100 &mu;m.

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No conflicts of interest declared.

This paper describes a versatile method for single cell fate mapping based on the photoactivation of caged fluorescein-dextran. Uncaging of caged fluorescein-dextran in single cells is achieved using two-photon absorption from a Mai Tai femtosecond laser coupled to the Zeiss LSM 510 META confocal microscope. Uncaged fluorescein-dextran in photoactivated cells is rapidly visualized using a simple immunohistochemistry procedure.
In this protocol the two-photon absorption wavelength for photoactivation of caged fluorescein-dextran is 740 nm. This wavelength was experimentally determined by photoactivating caged fluorescein-dextran injected wildtype embryos. The laser excitation wavelength was varied from 600 to 800 nm, and the presence or absence of fluorescein fluorescence post-activation was qualitatively determined. We found that 740 nm produced the strongest fluorescein signal following activation. Ideally, 740 nm should work for all model systems; however, we advise testing a range of wavelengths to determine the optimal two-photon excitation wavelength for your sample.
In caged fluorescein-dextran injected wildtype embryos, for each two-photon excitation wavelength tested we also varied the average laser power. For an excitation wavelength of 740 nm, an average laser power of 47 mW produced a stronger fluorescein signal at the activated region in comparison to lower average laser powers. Higher average laser powers did not produce stronger fluorescein signals, but induced ablation of the tissue. Since femtosecond laser pulses are efficient at ablating biological tissue15, we recommend determining the threshold average laser power for photoactivation that avoids tissue ablation.
Two-photon absorption occurs within a small interaction volume. To ensure proper activation of caged fluorescein-dextran, the cell must be brought into proper focus. In the above protocol, GFP positive cells were simultaneously imaged under white light to confirm cell focus. Therefore, white light acquisition in addition to other channels is advisable. After confirming cell focus, our protocol suggests magnifying the activated cell. A zoom factor of 50 to 70 is suggested, however, this value depends on the size of the chosen cell. Increasing the zoom isolates the activated cell from adjacent cells, and limits the scan area for two-photon activation. Ideally, the scan area should cover a large area of the cell.
Detection of single activated cells requires that the background staining be minimal. In the above immunodetection protocol it is important that the sample is blocked in blocking buffer for 5 to 6 hours (step 4.13). When developing with NBT/BCIP, uncaged fluorescein-dextran should become visible in 10 to 20 mins. If background staining is strong, we suggest a longer blocking time and additional washes to remove the antibody (step 4.17).
Figures 1 and 2 provide representative results of photoactivation and immunodetection. In Figure 1, early 1- to 2-cell stage wildtype embryos were injected with caged fluorescein-dextran. The embryos were raised to mid somitogenesis and mounted in low melting agarose in the lateral orientation. Figure 1(a,b) depict brightfield and fluorescent images of the embryo before photoactivation. Evidence that the embryo remained uncaged is demonstrated by the absence of fluorescein fluorescence in Figure 1(b). A small region within a somite was activated with a wavelength of 740 nm for 31 secs using an average laser power of 47 mW, Figure 1(c; arrow). Post-activation, two-photon uncaging of fluorescein-dextran occurred as shown by the fluorescence from the activated area, Figure 1(d; arrow). Activation was not accompanied by laser ablation, as the somitic tissue remained intact, Figure 1(c). Figure 2 demonstrates the immunodetection of uncaged fluorescein-dextran. A small region in the lateral plate mesoderm of a 10-somite stage embryo was photoactivated using the same laser parameters as mentioned in Figure 1. The embryo was raised and processed using steps 4.1 through 4.20. The arrow in Figure 2 locates the region activated, as indicated by the accumulation of blue precipitate. Only the activated location is clearly detected without interfering background staining.
Preparation of Solutions:
1 X PBT (1 L) 
100 mL 10 X PBS 
2 g BSA 
10 mL 20 % Tween
10 X PBS (1 L) 
80 g NaCl 
2 g KCl 
6.1 g Na2HPO4 
1.9 g KH2PO4 
ddH2O to 1 L 
pH to 7.3
4 % Paraformaldehyde (160 mL) 
32 % Paraformaldehyde (two 10 mL vials) 
8 mL 20 X PBS 
132 mL ddH2O
AP Buffer ( 50 mL) 
1 mL 5 M NaCl 
2.5 mL 1 M MgCl2 
5 mL 1 M Tris pH 9.5 
250 &mu;L 20 % Tween 
41.25 mL ddH2O
Antibody solution (for 1 sample) 
5.5 &mu;L Acetone powder 
40 &mu;L PBT 
0.8 &mu;L LS 
0.1 &mu;L Anti-fluorescein-AP antibody
2 % LS (360 &mu;L for 1 sample) 
7.2 &mu;L 100 % LS 
352.8 &mu;L PBT
1 X Danieau media (1 L)&Dagger; 
11.6 mL 5 M NaCl 
700 &mu;L 1 M KCl 
400 &mu;L 1 M MgSO4 
600 &mu;L 1 M Ca(NO3)2 
5 mL 1 M HEPES 
ddH2O to 1 L 
pH adjust to 7.6
&Dagger; Danieau is an ideal salt solution for rearing dechorionated zebrafish embryos. Other media may be used, provided they are isotonic solutions with properly osmolarity (~ 300 mOsm for zebrafish)
NBT stock (1 mL) 
50 mg Nitro Blue Tetrazolium 
0.7 mL dimethyl formamide anhydride 
0.3 mL ddH2O
BCIP stock (1 mL) 
50 mg 5-bromo-4-chloro-3-indolyl phosphate 
1 mL dimethyl formamide anhydride
NBT/BCIP developing solution 
1 mL AP buffer 
4.5 &mu;L NBT stock 
3.5 &mu;L BCIP stock
Acetone powder
<ol>
	<li>Select 20 adult fish and freeze them in liquid nitrogen.</li>
	<li>Grind the frozen fish with a mortar and pestle to a powder.</li>
	<li>Transfer the fish powder into a 50 mL conical tube.</li>
	<li>Fill the conical tube with 100 % acetone. Vortex the tube.</li>
	<li>Spin the tube at max rpm for 10 mins. Discard the supernatant.</li>
	<li>Wash the pellet another 3 times in 100 % acetone and spin the tube at max rpm for 10 mins. By the last wash the supernatant should appear clear.</li>
	<li>Add 100 % acetone to the pellet again and let the tube stand at room temperature. The more dense particles will settle, while the finer particles will remain in suspension.</li>
	<li>Decant the fine particles into a new tube. Aliquot the powder solution into separate tubes and store them at -20 &deg;C.</li>
</ol>

<table border="1">
	<tbody>
		<tr>
			<th>Name of the reagent</th>
			<th>Company</th>
			<th>Catalogue number</th>
		</tr>
		<tr>
			<td>CMNB-caged fluorescein SE</td>
			<td>Invitrogen</td>
			<td>C20050</td>
		</tr>
		<tr>
			<td>10 kDa Aminodextran</td>
			<td>Invitrogen</td>
			<td>D1860</td>
		</tr>
		<tr>
			<td>Zeba Desalt Spin Column		</td>
			<td>Pierce</td>
			<td>89889</td>
		</tr>
		<tr>
			<td>Low melting agarose</td>
			<td>Amresco</td>
			<td>0815-100G</td>
		</tr>
		<tr>
			<td>Sodium Borate</td>
			<td>Amresco</td>
			<td>1131117-100G</td>
		</tr>
		<tr>
			<td>Tricaine Methylsulfonate/ MS222</td>
			<td>Research Organics</td>
			<td>1347A</td>
		</tr>
		<tr>
			<td>Anti-fluorescein-AP</td>
			<td>Roche</td>
			<td>11376622</td>
		</tr>
		<tr>
			<td>Blocking buffer</td>
			<td>Roche Applied Sciences</td>
			<td>11 096 176 001</td>
		</tr>
		<tr>
			<td>Maleic Acid</td>
			<td>Sigma</td>
			<td>MO375-500G</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Sciences</td>
			<td>15714</td>
		</tr>
		<tr>
			<td>NBT	</td>
			<td>Sigma</td>
			<td>I8377-250mg</td>
		</tr>
		<tr>
			<td>BCIP</td>
			<td>Fischer Scientific</td>
			<td>PI-34040</td>
		</tr>
		<tr>
			<td>Microinjector</td>
			<td>Harvard Apparatus</td>
			<td>PLI-2000</td>
		</tr>
		<tr>
			<td>LabTek chambered coverglass slides</td>
			<td>Nalge Nunc International</td>
			<td>155380</td>
		</tr>
		<tr>
			<td>Proteinase K</td>
			<td>Invitrogen</td>
			<td>AM2544</td>
		</tr>
		<tr>
			<td>Lamb serum</td>
			<td>Invitrogen</td>
			<td>16070096</td>
		</tr>
		<tr>
			<td>LSM 510 META NLO</td>
			<td>Zeiss</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>20X 0.8 NA Air apochromatic objective</td>
			<td>Zeiss</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Transparent yellow filter</td>
			<td>Theatre House Inc.</td>
			<td>Roscolux filter sheet Color: 312</td>
		</tr>
	</tbody>
</table>

single cell fate mapping in zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

Watch this Scientific Journal Video about Single Cell Fate Mapping in Zebrafish at JoVE.com

Single Cell Fate Mapping in Zebrafish

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Research

JoVE Journal

Biology

13.4K ビュー.  Cincinnati Children's Hospital Medical Center. A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

ビデオ: Single Cell Fate Mapping in Zebrafish

Transplantation of Whole Kidney Marrow in Adult Zebrafish

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an organism. The functional definition of a HSC is a transplanted cell that has the ability to reconstitute all the blood lineages of an irradiated recipient long term. This designation was established by decades of seminal work in mammalian systems. Using hematopoietic cell transplantation (HCT) and reverse genetic manipulations in the mouse, the underlying regulatory factors of HSC biology are beginning to be unveiled, but are still largely under-explored. Recently, the zebrafish has emerged as a powerful genetic model to study vertebrate hematopoiesis. Establishing HCT in zebrafish will allow scientists to utilize the large-scale genetic and chemical screening methodologies available in zebrafish to reveal novel mechanisms underlying HSC regulation. In this article, we demonstrate a method to perform HCT in adult zebrafish. We show the dissection and preparation of zebrafish whole kidney marrow, the site of adult hematopoiesis in the zebrafish, and the introduction of these donor cells into the circulation of irradiated recipient fish via intracardiac injection. Additionally, we describe the post-transplant care of fish in an "ICU" to increase their long-term health. In general, gentle care of the fish before, during, and after the transplant is critical to increase the number of fish that will survive more than one month following the procedure, which is essential for assessment of long term (&lt;3 month) engraftment. The experimental data used to establish this protocol will be published elsewhere. The establishment of this protocol will allow for the merger of large-scale zebrafish genetics and transplant biology.

In this article, we demonstrate a method to perform HCT in adult zebrafish.

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing and adult tissue. An advance in this concept is Genetic Inducible Fate Mapping (GIFM), linking gene expression, cell fate, and cell behaviors in vivo, to create fate maps based on genetic lineage.
GIFM exploits X-CreER lines where X is a gene or set of gene regulatory elements that confers spatial expression of a modified bacteriophage protein, Cre recombinase (CreERT). CreERT contains a modified estrogen receptor ligand binding domain which renders CreERT sequestered in the cytoplasm in the absence of the drug tamoxifen. The binding of tamoxifen releases CreERT, which translocates to the nucleus and mediates recombination between DNA sequences flanked by loxP sites. In GIFM, recombination typically occurs between a loxP flanked Stop cassette preceding a reporter gene such as GFP.
Mice are bred to contain either a region- or cell type-specific CreER and a conditional reporter allele. Untreated mice will not have marking because the Stop cassette in the reporter prevents further transcription of the reporter gene. We administer tamoxifen by oral gavage to timed-pregnant females, which provides temporal control of CreERT release and subsequent translocation to the nucleus removing the Stop cassette from the reporter. Following recombination, the reporter allele is constitutively and heritably expressed. This series of events marks cells such that their genetic history is indelibly recorded. The recombined reporter thus serves as a high fidelity genetic lineage tracer that, once on, is uncoupled from the gene expression initially used to drive CreERT.
We apply GIFM in mouse to study normal development and ascertain the contribution of genetic lineages to adult cell types and tissues. We also use GIFM to follow cells on mutant genetic backgrounds to better understand complex phenotypes that mimic salient features of human genetic disorders.
This video article guides researchers through experimental methods to successfully apply GIFM. We demonstrate the method using our well characterized Wnt1-CreERT;mGFP mice by administering tamoxifen at embryonic day (E)8.5 via oral gavage followed by dissection at E12.5 and analysis by epifluorescence stereomicroscopy. We also demonstrate how to micro-dissect fate mapped domains for explant preparation or FACS analysis and dissect adult fate-mapped brains for whole mount fluorescent imaging. Collectively, these procedures allow researchers to address critical questions in developmental biology and disease models.

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Genetic Inducible Fate Mapping (GIFM) marks and tracks cells with fine spatial and temporal control in vivo and elucidates how cells from a specific genetic lineage contribute to developing and adult tissues. Demonstrated here are the techniques required to fate map E12.5 mouse embryos for epifluorescent and explant analysis.

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing ...

Fate Mapping of Human Embryonic Stem Cells by Teratoma Formation

Human embryonic stem cells (hESCs) have an unlimited capacity for self-renewal, and the ability to differentiate into cells derived from all three embryonic germ layers (1). Directed differentiation of hESCs into specific cell types has generated much interest in the field of regenerative medicine (e.g., (2-5)), and methods for determining the in vivo fate of selected or manipulated hESCs are essential to this endeavor. We have adapted a highly efficient teratoma formation assay for this purpose. A small number of specifically selected hESCs is mixed with undifferentiated wild type hESCs and Phaseolus vulgaris lectin to form a cell pellet. This is grafted beneath the kidney capsule in an immunodeficient mouse. As few as 2.5 x 105 hESCs are needed to form a 16 cm3 teratoma within 8-12 weeks. The fate of the originally selected hESCs can then be determined by immunohistochemistry. This method provides a valuable tool for characterizing tissue-specific reagents for cell-based therapy.

Directed differentiation of hESCs into specific cells has generated much interest in regenerative medicine. We provide a concise, step-by-step protocol for determining the in vivo fate of selected hESCs that provides a valuable tool for characterizing tissue-specific reagents for cell-based therapy.

Human embryonic stem cells (hESCs) have an unlimited capacity for self-renewal, and the ability to differentiate into cells derived from all three ...

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal neurons. In contrast, the adult zebrafish retina possesses the robust ability to spontaneously regenerate any neuronal class that is lost in a variety of different retinal damage models, including retinal puncture, chemical ablation, concentrated high temperature, and intense light treatment 1-8. Our lab extensively characterized regeneration of photoreceptors following constant intense light treatment and inner retinal neurons after intravitreal ouabain injection 2, 5, 9. In all cases, resident M&uuml;ller glia re-enter the cell cycle to produce neuronal progenitors, which continue to proliferate and migrate to the proper retinal layer, where they differentiate into the deficient neurons. 
	We characterized five different stages during regeneration of the light-damaged retina that were highlighted by specific cellular responses. We identified several differentially expressed genes at each stage of retinal regeneration by mRNA microarray analysis 10. Many of these genes are also critical for ocular development. To test the role of each candidate gene/protein during retinal regeneration, we needed to develop a method to conditionally limit the expression of a candidate protein only at times during regeneration of the adult retina. 
	Morpholino oligos are widely used to study loss of function of specific proteins during the development of zebrafish, Xenopus, chick, mouse, and tumors in human xenografts 11-14. These modified oligos basepair with complementary RNA sequence to either block the splicing or translation of the target RNA. Morpholinos are stable in the cell and can eliminate or "knockdown" protein expression for three to five days 12. 
	Here, we describe a method to efficiently knockdown target protein expression in the adult zebrafish retina. This method employs lissamine-tagged antisense morpholinos that are injected into the vitreous of the adult zebrafish eye. Using electrode forceps, the morpholino is then electroporated into all the cell types of the dorsal and central retina. Lissamine provides the charge on the morpholino for electroporation and can be visualized to assess the presence of the morpholino in the retinal cells. 
	Conditional knockdown in the retina can be used to examine the role of specific proteins at different times during regeneration. Additionally, this approach can be used to study the role of specific proteins in the undamaged retina, in such processes as visual transduction and visual processing in second order neurons.

In vivo Electroporation of Morpholinos into the Adult Zebrafish Retina

A method to conditionally knockdown a target protein&#x2019;s expression in the adult zebrafish retina is described, which involves intravitreally injecting antisense morpholinos and electroporating them into the retina. The resulting protein is knocked down for several days, which allows testing the protein&#x2019;s role in the regenerating or intact retina.

Many devastating inherited eye diseases result in progressive and irreversible blindness because humans cannot regenerate dying or diseased retinal ...

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration of adult tissues, such as the heart1, retina2, spinal cord3, optic nerve4, sensory hair cells5, and fins6.
The zebrafish fin is a relatively simple appendage that is easily manipulated to study multiple stages in epimorphic regeneration. Classically, fin regeneration was characterized by three distinct stages: wound healing, blastema formation, and fin outgrowth. After amputating part of the fin, the surrounding epithelium proliferates and migrates over the wound. At 33 &deg;C, this process occurs within six hours post-amputation (hpa, Figure 1B)6,7. Next, underlying cells from different lineages (ex. bone, blood, glia, fibroblast) re-enter the cell cycle to form a proliferative blastema, while the overlying epidermis continues to proliferate (Figure 1D)8. Outgrowth occurs as cells proximal to the blastema re-differentiate into their respective lineages to form new tissue (Figure 1E)8. Depending on the level of the amputation, full regeneration is completed in a week to a month.
The expression of a large number of gene families, including wnt, hox, fgf, msx, retinoic acid, shh, notch, bmp, and activin-betaA genes, is up-regulated during specific stages of fin regeneration9-16. However, the roles of these genes and their encoded proteins during regeneration have been difficult to assess, unless a specific inhibitor for the protein exists13, a temperature-sensitive mutant exists or a transgenic animal (either overexpressing the wild-type protein or a dominant-negative protein) was generated7,12. We developed a reverse genetic technique to quickly and easily test the function of any gene during fin regeneration.
Morpholino oligonucleotides are widely used to study loss of specific proteins during zebrafish, Xenopus, chick, and mouse development17-19. Morpholinos basepair with a complementary RNA sequence to either block pre-mRNA splicing or mRNA translation. We describe a method to efficiently introduce fluorescein-tagged antisense morpholinos into regenerating zebrafish fins to knockdown expression of the target protein. The morpholino is micro-injected into each blastema of the regenerating zebrafish tail fin and electroporated into the surrounding cells. Fluorescein provides the charge to electroporate the morpholino and to visualize the morpholino in the fin tissue.
This protocol permits conditional protein knockdown to examine the role of specific proteins during regenerative fin outgrowth. In the Discussion, we describe how this approach can be adapted to study the role of specific proteins during wound healing or blastema formation, as well as a potential marker of cell migration during blastema formation.

In vivo Electroporation of Morpholinos into the Regenerating Adult Zebrafish Tail Fin

We describe a method to conditionally knockdown the expression of a target protein during adult zebrafish fin regeneration. This technique involves micro-injecting and electroporating antisense oligonucleotide morpholinos into fin tissue, which allows testing the protein&#x2019;s role in various stages of fin regeneration, including wound healing, blastema formation, and regenerative outgrowth.

Certain species of urodeles and teleost fish can regenerate their tissues. Zebrafish have become a widely used model to study the spontaneous regeneration ...

Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal which tissues descend from each cell of the embryo. Although fate maps are very useful for identifying the precursors of an organ and for elucidating the developmental path by which the descendant cells populate that organ in the normal embryo, they do not illustrate the full developmental potential of a precursor cell or identify the mechanisms by which its fate is determined. To test for cell fate commitment, one compares a cell's normal repertoire of descendants in the intact embryo (the fate map) with those expressed after an experimental manipulation. Is the cell's fate fixed (committed) regardless of the surrounding cellular environment, or is it influenced by external factors provided by its neighbors? Using the comprehensive fate maps of the Xenopus embryo, we describe how to identify, isolate and culture single cleavage stage precursors, called blastomeres. This approach allows one to assess whether these early cells are committed to the fate they acquire in their normal environment in the intact embryo, require interactions with their neighboring cells, or can be influenced to express alternate fates if exposed to other types of signals.

The fate of an individual embryonic cell can be influenced by inherited molecules and/or by signals from neighboring cells. Utilizing fate maps of the cleavage stage Xenopus embryo, single blastomeres can be identified for culture in isolation to assess the contributions of inherited molecules versus cell-cell interactions.

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal  which tissues descend from each cell of the embryo.  Although fate maps ...

Generation of Dispersed Presomitic Mesoderm Cell Cultures for Imaging of the Zebrafish Segmentation Clock in Single Cells

Segmentation is a periodic and sequential morphogenetic process in vertebrates. This rhythmic formation of blocks of tissue called somites along the body axis is evidence of a genetic oscillator patterning the developing embryo. In zebrafish, the intracellular clock driving segmentation is comprised of members of the Her/Hes transcription factor family organized into negative feedback loops. We have recently generated transgenic fluorescent reporter lines for the cyclic gene her1 that recapitulate the spatio-temporal pattern of oscillations in the presomitic mesoderm (PSM). Using these lines, we developed an in vitro culture system that allows real-time analysis of segmentation clock oscillations within single, isolated PSM cells. By removing PSM tissue from transgenic embryos and then dispersing cells from oscillating regions onto glass-bottom dishes, we generated cultures suitable for time-lapse imaging of fluorescence signal from individual clock cells. This approach provides an experimental and conceptual framework for direct manipulation of the segmentation clock with unprecedented single-cell resolution, allowing its cell-autonomous and tissue-level properties to be distinguished and dissected.

Somitogenesis is a rhythmic developmental process that spatially patterns the body axis of vertebrate embryos. Previously, we developed transgenic zebrafish lines that use fluorescent reporters to observe the cyclic genes that drive this process. Here, we culture dispersed cells from these lines and image their oscillations over time in vitro.

Segmentation is a  periodic and sequential morphogenetic process in vertebrates. This rhythmic  formation of blocks of tissue called somites along the ...

Identification of MyoD Interactome Using Tandem Affinity Purification Coupled to Mass Spectrometry

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ability to proliferate or they can differentiate and fuse into multinucleated myotubes, which maturate further to form myofibers. Skeletal muscle terminal differentiation is orchestrated by the coordinated action of various transcription factors, in particular the members of the Muscle Regulatory Factors or MRFs (MyoD, Myogenin, Myf5, and MRF4), also called the myogenic bHLH transcription factors family. These factors cooperate with chromatin-remodeling complexes within elaborate transcriptional regulatory network to achieve skeletal myogenesis. In this, MyoD is considered the master myogenic transcription factor in triggering muscle terminal differentiation. This notion is strengthened by the ability of MyoD to convert non-muscle cells into skeletal muscle cells. Here we describe an approach used to identify MyoD protein partners in an exhaustive manner in order to elucidate the different factors involved in skeletal muscle terminal differentiation. The long-term aim is to understand the epigenetic mechanisms involved in the regulation of skeletal muscle genes, i.e., MyoD targets. MyoD partners are identified by using Tandem Affinity Purification (TAP-Tag) from a heterologous system coupled to mass spectrometry (MS) characterization, followed by validation of the role of relevant partners during skeletal muscle terminal differentiation. Aberrant forms of myogenic factors, or their aberrant regulation, are associated with a number of muscle disorders: congenital myasthenia, myotonic dystrophy, rhabdomyosarcoma and defects in muscle regeneration. As such, myogenic factors provide a pool of potential therapeutic targets in muscle disorders, both with regard to mechanisms that cause disease itself and regenerative mechanisms that can improve disease treatment. Thus, the detailed understanding of the intermolecular interactions and the genetic programs controlled by the myogenic factors is essential for the rational design of efficient therapies.

MyoD is a myogenic transcription factor with a strong capacity to induce myogenic transdifferentiation of many fully differentiated non-muscle cell lines. The epigenetic mechanisms involved in this transdifferentiation are largely unknown. Here we describe a double-affinity purification method followed by mass spectrometry to exhaustively characterize MyoD partners.

Skeletal muscle terminal differentiation starts with the commitment of pluripotent mesodermal precursor cells to myoblasts. These cells have still the ...

Open-source Single-particle Analysis for Super-resolution Microscopy with VirusMapper

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm allows for the routine imaging of nanoscale biological complexes and processes. This increase in resolution also means that methods popular in electron microscopy, such as single-particle analysis, can readily be applied to super-resolution fluorescence microscopy. By combining this analytical approach with super-resolution optical imaging, it becomes possible to take advantage of the molecule-specific labeling capacity of fluorescence microscopy to generate structural maps of molecular elements within a metastable structure. To this end, we have developed a novel algorithm &#8212; VirusMapper &#8212; packaged as an easy-to-use, high-performance, and high-throughput ImageJ plugin. This article presents an in-depth guide to this software, showcasing its ability to uncover novel structural features in biological molecular complexes. Here, we present how to assemble compatible data and provide a step-by-step protocol on how to use this algorithm to apply single-particle analysis to super-resolution images.

This manuscript uses the Fiji-based open-source software package VirusMapper to apply single-particle analysis to super-resolution microscopy images in order to generate precise models of nanoscale structure.

Super-resolution fluorescence microscopy is currently revolutionizing cell biology research. Its capacity to break the resolution limit of around 300 nm ...