PLATIM and Droso-Tools facilities of the UMS3444 Biosciences, Lyon, France. BM&#39;s research was supported by grants from the Fondation pour la Recherche M&eacute;dicale from the CNRS (ATIP) and the ANR-12-BSV1-0019-01. PD was supported by Retina France association and the Ecole Normale Sup&eacute;rieure of Lyon (France).&nbsp;CL was supported by La Ligue Nationale contre le Cancer association.&nbsp;

1. Crossing with Tomato/GFP-FLP/FRT Lines

	The generation of flies carrying the mutant clone requires a single cross between a FRT-carrying mutant line and the corresponding Tomato/GFP-FLP/FRT fly line.

	This work utilized the BruinFly collection of FRT-recombined mutations (<a href="http://www.bruinfly.ucla.edu" target="_blank">http://www.bruinfly.ucla.edu</a>). Four Tomato/GFP-FLP/FRT lines can be used, carrying FRT Rh1-tdtomatoninaC on each arms of the 2nd and 3rd chromosomes combined with source of flipase (ey-FLP) and expression of GFP in all outer PRs (Rh1-Gal4 UAS-GFP) 20:

	- 2L arm: line #43345, P{ry[+t7.2]=rh1-GAL4}1, P{ry[+t7.2]=ey-FLP.N}2, w[*]; P{w[+mC]=ninaE-tdTomato-ninaC}2L P{ry[+t7.2]=neoFRT}40A; P{w[+mC]=UAS-GFP-ninaC}3

	- 2R arm: line #43346, P{ry[+t7.2]=rh1-GAL4}1, P{ry[+t7.2]=ey-FLP.N}2, w[*]; P{ry[+t7.2]=neoFRT}42D P{w[+mC]=ninaE-tdTomato-ninaC}2R; P{w[+mC]=UAS-GFP-ninaC}3

	- 3L arm: line #43347, P{ry[+t7.2]=rh1-GAL4}1, P{ry[+t7.2]=ey-FLP.N}2, w[*]; P{w[+mC]=UAS-GFP-ninaC}2; P{w[+mC]=ninaE-tdTomato-ninaC}3L P{ry[+t7.2]=neoFRT}80B/TM6B, Tb[1]

	- 3R arm: line #43348, P{ry[+t7.2]=rh1-GAL4}1, P{ry[+t7.2]=ey-FLP.N}2, w[*]; P{w[+mC]=UAS-GFP-ninaC}2; P{ry[+t7.2]=neoFRT}82B P{w[+mC]=ninaE-tdTomato-ninaC}3R/TM6B, Tb[1]

	These four lines are available at the Bloomington Drosophila stock center. They provided a source of FLP in the developing eye for the generation of mitotic clones 21. The wild-type chromosome carries an FRT sequence and a rh1-tdTomatoninaC construct encoding the red fluorescent protein tdTomato, as a marker for wild-type and heterozygous outer PRs. In this construct, the last 41 amino acids of the p174 isoform of the ninaC (neither inactivation nor activation C) gene was appended in-frame to the C-terminus of the tdTomato sequence. The C-terminal tail of the p174 ninaC isoform is responsible for the rhabdomeral localization of the protein 22 and permits better live visualization of red fluorescent PRs using the cornea neutralization technique. The rh1 promoter is the minimal 234 bp (-152+82) rh1 promoter. The green fluorescent marker GFP is expressed by all the outer PRs, due to the presence of the rh1-Gal4 (3kb promoter) and UAS-GFPninaC constructs. Crosses between a fly stock carrying a mutation x on the FRT chromosome (FRT-x) and the Tomato/GFP-FLP/FRT lines yields a progeny with the following genotype (example of mutation on 2L): rh1-Gal4, ey-FLP; FRT40A, rh1-tdTomatoninaC/FRT40A-x; UAS-GFPninaC. In these flies, homozygous mutant cells are generated by mitotic recombination in the developing eye disc and divide to generate a clone of homozygous mutant PRs (Figure 2A). Homozygous mutant cells can be identified because they express only the green fluorescent protein (GFP), whereas the surrounding wild-type and heterozygous cells express both tdTomato and GFP (Figures 2B and B").

	2. Preparing Drosophila for Photoreceptor Visualization

	For PR visualization by the cornea neutralization technique, immobilize flies onto agarose plates.
<ol>
	<li>
		Prepare a wash bottle filled with 4 &deg;C distilled water and place it on ice.</li>
	<li>
		Dissolve regular agarose in water (1.2-1.5% agarose in 100 ml of water) by heating in a microwave. Place the flask in a water bath at 55 &deg;C and allow the agarose solution to cool to this temperature. Keep the agarose at 55 &deg;C until use.</li>
	<li>
		Anesthetize the flies with CO2 for at least 1 min.</li>
	<li>
		Pour the warm agarose solution (at 55 &deg;C) into a Petri dish (35 x 10 mm, 1.37 x 0.39 in) and place immediately the anesthetized Drosophila onto the agarose. For beginners, starting with 10 flies per Petri dish is reasonable. Bigger Petri dish (60 x 15 mm, 2.362 x 0.59 in) can also be used.</li>
	<li>
		Under a dissecting microscope, orient the Drosophila on its side with forceps, push one wing into the agarose, such that the wing and half of the body are embedded in the agarose (Figures 3A-3A""). Stick the other wing onto the surface of the agarose. Note: If the flies are not embedded deeply enough in the agarose, the fly will be free to move its head during imaging of the PR, and this will result in fuzzy images. The orientation of the eye will also be disturbed. It may be difficult to plunge the Drosophila into the agarose, due to either the concentration of the agarose or its temperature. It is easier to embed flies in more concentrated agarose, but if the agarose is too concentrated, the fly may sink. If the agarose is too cool, it may be difficult to embed the fly, but too high a temperature may damage the cornea.</li>
	<li>
		Put the Petri dish on ice and allow the agarose to solidify.</li>
	<li>
		With forceps, orient the head such that one eye is exposed to the immersion objective (Figure 3A' and 3A""). Generally, an eye may be considered to be well oriented under the dissecting microscope when the pseudopupil (visualized as a black spot) is in the middle of the eye. An optimal orientation of the eye maximizes the width of the ommatidial field in which PRs are focused. The aim of the orientation step is to find the region of the eye with the widest field of focused PRs, usually in the center of the eye. This step is also useful for removing any legs or aristae covering the eye and preventing its visualization.</li>
	<li>
		Cover the fly with ice-cold water and leave the Petri dish on ice until visualization. Ice-cold water keeps the flies anesthetized.</li>
</ol>

	3. Visualizing Photoreceptors under the Microscope

	For the visualization of PRs through the cornea, this protocol uses an upright microscope equipped with a water objective with a long working distance (W N-Achroplan 40X/0.75).
<ol>
	<li>
		Place the Petri dish covered with ice-cold water on a glass slide on the stage of the microscope (Figures 3 B-C). The Petri dish can be glued to the glass slide, making it possible to move it about smoothly with the micrometer screws.</li>
	<li>
		Plunge the immersion objective into the water of the Petri dish (Figures 3B'-3C'). Position the head of the fly under the excitation beam (start with the GFP filter, for example). Move the stage up and down until the beam converges on the eye. When the eye is at the right level, it tends to reflect the excitation light.</li>
	<li>
		Look through the eyepiece for fluorescence. When the eye is positioned in the center of the field of view, focus below the cornea to visualize the fluorescent photoreceptors.</li>
</ol>

	Note 1: When looking through the eyepiece, it may be uneasy to distinguish the body parts of the fly, particularly the distal part of the abdomen and the head, for example. In such cases, you should look directly at the stage to reposition the Drosophila.

	Note 2: Classical fluorescence or confocal microscopy may be used. Confocal microscopy gives images with less background and a wider field of focused PRs than classical microscopy. An example set-up is an LSM510 confocal microscope (Zeiss) with a 40X water objective. Better results may be obtained by opening the pinhole of the confocal microscope wider than is usually recommended for the objective. For example, use a value of 204, rather than the default 98 for the aperture of the pinhole.

	Note 3: Higher levels of w+ pigmentation of the eye reduce the background fluorescence.

	4. Time-course Visualization of Photoreceptor Cells

	Care is required when following the fate of an individual PR within the same eye over a period of time.
<ol>
	<li>
		Reduce the temperature of the agarose to 45 &deg;C, to maximize Drosophila survival. The agarose solidifies very quickly at this temperature. Only one Drosophila should be used per Petri dish, to ensure that flies are not mixed up during observations.</li>
	<li>
		Systematically position the flies on the same side, for viewing of the same eye. This can be achieved by taking the anesthetized fly from its vial by one wing and placing the Drosophila on the agarose. Always orient the eye in the same way if possible, making it easy to find PR clones that have been viewed before.</li>
	<li>
		Identify the clones on the basis of their shape under the confocal microscope. The field of view is often located next to the clone of interest. In such cases, the eye should be reoriented under water, to center the field of view on the clone of interest, based on the polarity of the eye (Figure 4).</li>
</ol>

	5. Recovering Flies after Visualization
<ol>
	<li>
		Remove the water from the Petri dish.</li>
	<li>
		Gently pull the Drosophila out of the agarose with forceps.</li>
	<li>
		Dry the Drosophila on a tissue.</li>
	<li>
		Return the Drosophila to a vial, ensuring that it does not get stuck in the food. Allowed the fly to come round at 25 &deg;C.</li>
</ol>

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The authors have nothing to disclose.

The Drosophila eye has been widely used to decipher the signaling pathways regulating development, proliferation and survival. In the early 1990s, a large number of genetic screens were carried out to identify the pathways required during the initial phases of PR development 25-27. The efficiency of genetic screens was increased greatly by the mitotic recombination technique, which makes it possible to test loss-of-function mutations in mosaic clones 21. Hence, the role of embryonic lethal mutations could be systematically tested in homozygous mutant clones in the eyes of flies that were otherwise heterozygous. Most mosaic screenings have focused on the early PR recruitment, differentiation, axonal projection or morphogenesis occurring in the developing larvae or pupae 10,25-31. To date, only one mosaic screen has investigated the mechanisms regulating adult PR function, by monitoring the visual response via electroretinogram recordings 32. With the Tomato/GFP-FLP/FRT method and the possibility of performing time course analysis in living mutant animals, we have designed a new method for identifying the factors regulating adult PR viability and function 20,24.

	The Tomato/GFP-FLP/FRT method has several major advantages over the classical histological sectioning of resin-embedded eyes. First, this method is much faster, cheaper and easier to perform than the histological method, provided that a fluorescence microscope equipped with an appropriate water dipping objective is available (see Table of Reagents and Equipment). Second, the Tomato/GFP-FLP/FRT method can be used in conjunction with transgene expression, such as the expression of P(UAS, w+), carrying a mini-white transgene. In contrast to histological sections, in which w+ is used for clonal detection, in the Tomato/GFP-FLP/FRT system, P(UAS, w+) does not interfere with clonal detection based on Tomato fluorescent protein. Using P(UAS-fatp, w+) transgenic flies, we were able to visualize the rescue of fatp mutant PRs (Figure 7). Third, a key pitfall of all types of clonal analysis, including that based on Tomato/GFP-FLP/FRT, is the ambiguity of the genotype of lost PRs. Indeed, it may be unclear whether a PR is absent because of the lack of a specific gene function or because of a cell non-autonomous effect, particularly at the border of the mutant clone. The Tomato/GFP-FLP/FRT method gets around this problem for degeneration by making it possible to follow wild-type and mutant PRs in the same animal over periods of several weeks. We were able to follow the fate of individual PRs by kinetic analyses on different fatp mutant flies (Figure 6). The same clone can be recognized in a given animal at different times, because of the precise shape of the surrounding wild-type clones. We were able to show unambiguously that all the missing PRs were mutant for fatp, indicating that the role of fatp mutants in PRs is cell-autonomous. Thus, by monitoring the loss of PRs in a kinetic analysis, it is possible to determine the genotype of PRs in models of adult-onset degeneration. Finally, the Tomato/GFP-FLP/FRT method has proved very powerful for the identification of factors regulating the establishment of PCP 20. The possibility of scoring a large number of mosaic ommatidia for a polarity phenotype without the need for sections, facilitates rapid determination of the PR requirement for the establishment of PCP. Nevertheless, finer analysis of PR integrity would require traditional sectioning followed by phase-contrast or electronic microscopies.

	In conclusion, the Tomato/GFP-FLP/FRT method opens up new possibilities for efficient mosaic screening to identify factors regulating PR developmental processes and adult PR functions, such as the visual response and viability.

The Drosophila retina is composed of about 800 ommatidial units (Figure 1A), which are precisely organized, with a clearly defined axis of polarity (Figure 1B). Each unit contains 20 cells: eight photoreceptor cells (PRs) and 12 accessory cells, including cone cells, pigment cells and bristle cells 1,2. Two classes of PR are distinguished on the basis of the type of rhodopsin (Rh) they express. The six outer PRs (R1-6) express Rh1 and are arranged in a trapezoid pattern (Figure 1C). In the center of the trapezoid, the two inner PRs (R7/R8) express four possible types of rhodopsin (Rh3, Rh4, Rh5 or Rh6) and are organized such that R7 lies on top of R8 3.

	More than 2,500 genes are involved in the morphogenesis of the Drosophila eye 4, which has proved a very powerful model for studies of a wide panel of processes, including eye development, PR recruitment, differentiation, planar cell polarity, morphogenesis, survival, apoptosis and visual transduction 5-9.

	Researchers working on the Drosophila eye have, over many years, developed techniques for imaging the retina and carrying out systematic genetic screens. The easiest way to image the adult retina is to look at the cornea in an immobilized animal (Figure 1A). The structure of the cornea can be visualized precisely by scanning electron microscopy and was used in a large-scale genetic screen by the URCFG consortium (<a href="http://www.bruinfly.ucla.edu" target="_blank">http://www.bruinfly.ucla.edu</a>)10. This is a very effective approach for visualizing global morphological changes in corneal structure, such as eye roughness or glossiness, often induced by mutations in genes controlling early steps in development or cell viability. However, global visualization of the cornea is not sufficient to identify the factors regulating PR rhabdomere morphogenesis or adult PR viability and function. Such analyses require a more thorough investigation of PR integrity, based on phase-contrast microscopy of tangential semi-thin resin sections of the retina 11-13. This technique is suitable for mosaic analyses, in which mutant PRs can be identified by their lack of red pigment 14,15.
Mosaic mutant clones can also be visualized in whole-mount dissections of the pupal or adult retina, on the basis of their lack of green fluorescent protein (GFP) signal on fluorescence microscopy 16,17. These two techniques are very useful, but both are labor- and time-consuming and are therefore not suitable for large-scale screening. We and others have developed the use of cornea neutralization techniques for the imaging of PRs producing fluorescent proteins 18,19, to facilitate rapid PR analyses. With this technique, mutant clones can be identified on the basis of the autofluorescence of the red (w+) pigment in mosaic adult Drosophila clones. However, this method does not provide the single-cell resolution required to address cell autonomy issues 19. We overcame this problem by developing the Tomato/GFP-FLP/FRT method, which combines the imaging of fluorescent proteins by cornea neutralization with mitotic recombination 20. This method allows the high-throughput, rapid and precise identification of mutant PRs in mosaic clones, at single-cell resolution (<a href="http://www.ens-lyon.fr/LBMC/ApoDrosoDatabase/" target="_blank">http://www.ens-lyon.fr/LBMC/ApoDrosoDatabase/</a>). It is suitable for use in kinetic analyses, for following individual PRs over a period of weeks in living Drosophila. We describe here the Tomato/GFP-FLP/FRT method and tips for its use in simple, kinetic analyses of mosaic eyes.

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				Regular agarose is used to immobilize the flies</td>
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				Dissecting microscope</td>
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				Dutcher</td>
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				SMZ645</td>
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				To keep the agarose solution at warm temperature</td>
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				420967-9900-000</td>
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				Water dipping objective for the confocal microscope</td>
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	&nbsp;

the tomato/gfp-flp/frt method for live imaging of mosaic adult drosophila photoreceptor cells

The Drosophila eye is widely used as a model for studies of development and neuronal degeneration. With the powerful mitotic recombination technique, elegant genetic screens based on clonal analysis have led to the identification of signaling pathways involved in eye development and photoreceptor (PR) differentiation at larval stages. We describe here the Tomato/GFP-FLP/FRT method, which can be used for rapid clonal analysis in the eye of living adult Drosophila. Fluorescent photoreceptor cells are imaged with the cornea neutralization technique, on retinas with mosaic clones generated by flipase-mediated recombination. This method has several major advantages over classical histological sectioning of the retina: it can be used for high-throughput screening and has proved an effective method for identifying the factors regulating PR survival and function. It can be used for kinetic analyses of PR degeneration in the same living animal over several weeks, to demonstrate the requirement for specific genes for PR survival or function in the adult fly. This method is also useful for addressing cell autonomy issues in developmental mutants, such as those in which the establishment of planar cell polarity is affected.

The Tomato/GFP-FLP/FRT method can be used to study the effect of a mutation or of ectopic expression on the development and survival of PRs in the Drosophila retina. It is rapid, making it ideal for screening purposes, as recently demonstrated 20. The presence of mutant PRs next to wild-type PRs in the same eye makes it easy to detect defects associated with mutant PRs and to address the cell autonomy issue.

 The developmental defects observed in our analysis affected various processes, including PR recruitment, morphogenesis and planar cell polarity (PCP) establishment (Figure 5). Seven in absentia is required for PR recruitment, particularly for R7 one of the inner PR. The loss of Seven in absentia results in the loss of the inner PR and some outer PRs (Figure 5A). Grainy head (grh) is involved in PCP establishment and some ommatidia are inverted in grh mutant clones (Figure 5B). This method makes it possible to identify the mutant PRs in mosaic ommatidia, facilitating identification of the PRs in which the gene is required for the correct acquisition of PCP. We have shown that grh is required for the correct acquisition of PCP in R3 precursors 20. Indeed, we observed that, in the inverted ommatidia, R4, which originates abnormally from the R3 precursor, was always mutant. Crumbs (Crbs) is an apical membrane protein required for rhabdomere morphogenesis 23. The loss of Crbs in the Crbs11A22 mutant results in irregular, larger or smaller PRs (Figure 5C).

 This method can also be used to follow the fate of single PRs during adulthood (Figure 6). It is ideal for neurodegeneration studies, because a group of homozygous mutant PRs can be viewed and the same group can be found in the same eye of the same fly over a period of days or weeks. It is therefore possible to determine which PRs survive and which are doomed to die, because each clone has a unique shape that can be recognized when the eye is positioned under the fluorescence microscope. As the retina is polarized, it is possible to orient the retina to find the same clone again, on the basis of its shape. We used this method to show that fatpk10307 mutation induces progressive PR degeneration in adulthood (Figure 6, 24). This visualization method can also be used to analyze cell autonomy. In fatpk10307 mosaic retina, PR loss was restricted to fatp mutant PRs, the wild-type PRs being unaffected. The requirement of fatp for PR viability is therefore cell autonomous. We were also able to rescue the fatp mutant PR by reexpressing wild-type fatp with a rh1-Gal4 driver and a UAS-fatp construct (Figure 7). The possibility of carrying out such rescue experiments is one of the advantages of the Tomato/GFP-FLP/FRT method over clonal analysis based on histological sections of resin-embedded retina. Indeed, clonal detection with the Tomato/GFP-FLP/FRT method is not affected by the use of P(UAS, w+) transgenic constructs, whereas the red pigmentation associated with the mini-white gene (w+) covers the eye with red pigment, masking mosaic clones in histological sections.

 <img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50610/50610fig1.jpg" /> 
 Figure 1. Organization of the Drosophila eye. (A) Photograph of a Drosophila eye taken with a stereomicroscope. The Drosophila eye is composed of approximately 800 ommatidia. (B) Schematic diagram of a field of 64 ommatidia in the middle of the eye. The six outer photoreceptor neurons (PRs) of each ommatidium, shown in green, are organized into a stereotypical trapezoid that points to the nearest poles of the eye. They consequently point in opposite directions in the ventral and the dorsal part of the eye and are mirror image according to the border between the two parts, which is called the equator. (C) Schematic diagram of an ommatidium. Each ommatidium contains eight PRs. PRs R1 to R6, shown in green, are the outer PRs. They express the photosensitive molecule rhodopsin1 (Rh1) and are the equivalent of mammalian rod cells. The inner PRs, R7 and R8, are shown in gray (R7 sits on top of R8). The inner PRs express rhodopsin 3, 4, 5 or 6 and are the equivalent of mammalian cone cells. For simplicity, only the rhabdomere, the light-sensitive part of the PR, is shown for each PR. <a href="https://www.jove.com/files/ftp_upload/50610/50610fig1large.jpg" target="_blank"> Click here to view larger figure</a>.

 <img alt="Figure 2" fo:content-width="4.25in" src="/files/ftp_upload/50610/50610fig2.jpg" /> 
 Figure 2. Generating and visualizing mosaic PR clones in the Drosophila eye with the Tomato/GFP-FLP/FRT method. (A) Schematic diagram of the generation of mosaic clones in the Tomato/GFP-FLP/FRT method. Mosaic clones are generated by mitotic recombination, due to the expression of the flipase (under control of the eyeless promoter) during eye development and the FRT sequences. After recombination of the FRT sequence and cell division, homozygous mutant, homozygous wild-type and heterozygous cells can be generated from a heterozygous cell. These cells divide again to form mosaic clones of PRs in the adult eye. The identification of mutant cells is facilitated by inserting a construct expressing the red fluorescent protein tdTomato into the wild-type chromosome, to label wild-type and heterozygous cells. (B-B'') Visualization of mosaic PR clones with the Tomato/GFP-FLP/FRT method. The Tomato/GFP-FLP/FRT method combines mitotic recombination, cornea neutralization and confocal microscopy. All PRs express the green fluorescent protein GFP, which facilitate the visualization of the PRs using cornea neutralization and confocal microscopy (B). PRs can be visualized at a single-cell level. Mutant mosaic clones are generated by mitotic recombination and can be identified by the absence of the red fluorescent protein tdTomato (B&#39;). Consequently, on merged images, homozygous PRs appear in green, whereas wild-type and heterozygous PRs are yellow (B''). <a href="https://www.jove.com/files/ftp_upload/50610/50610fig2large.jpg" target="_blank"> Click here to view larger figure</a>.

 <img alt="Figure 3" fo:content-width="6in" src="/files/ftp_upload/50610/50610fig3.jpg" /> 
 Figure 3. Setting up the Drosophila for visualization by cornea neutralization. (A-A'''') Photographs of a Drosophila immobilized on a plate filled with agarose. In panel A'''', a piece of agarose containing the fly has been cut to take a profile photograph. The Drosophila is half-embedded in the agarose, with the right wing within the agarose and the left wing stuck on the agarose surface (A, A'', A''''). When the Drosophila is embedded in the agarose, its head is frequently poorly positioned for the visualization of the eye (A, A'). The head must therefore be reorientated with forceps such that the middle of the eye is pointing upwards (A'', A'''). (B-B') Schematic diagram showing a Drosophila immobilized on an agarose plate and the positioning of the Drosophila under the objective of the microscope. (C-C') Photographs of the setting of the Drosophila on the stage of the microscope under its objective. <a href="https://www.jove.com/files/ftp_upload/50610/50610fig3large.jpg" target="_blank"> Click here to view larger figure</a>.

 <img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/50610/50610fig4.jpg" /> 
 Figure 4. Orientation of the eye for time-course analysis. The figure shows photographs of a Drosophila head in two different orientations and the corresponding fields of ommatidia viewed by cornea neutralization, represented as schematic diagrams. (A) During the first observation, the eye is placed such that a clone of heterozygous or wild-type PRs (in yellow) is viewed in the middle of the field, at the level of the equator (red line). (B) During the second observation, the eye is often not in exactly the same orientation. Thus, the same group of PRs can be found again, but it is not possible to view the exact same field (on the left). The eye must be reoriented for the same field to be viewed again. The position of the equator can be used as a guide. In this example, we know that the observed field is too ventral and that the eye must therefore be turned ventrally to place the equator in the middle of the observed field again, as in the first observation.

 <img alt="Figure 5" fo:content-width="6in" src="/files/ftp_upload/50610/50610fig5.jpg" /> 
 Figure 5. Examples of PR development defects detected by the Tomato/GFP-FLP/FRT method. (A) Visualization of mosaic sina[BG02648] mutant PRs, showing the loss of inner PRs. In the mutant ommatidia, outer PRs are clustered together due to the absence of the inner PR R7. Indeed, sina is known to be required for inner PR recruitment. (B) Visualization of mosaic grh[s2140] mutant PRs showing polarity defects. The mosaic mutant ommatidia, surrounded by a circle, are pointing in the opposite direction from wild-type ommatidia, indicating a dorso-ventral inversion. A detailed study with the Tomato/GFP-FLP/FRT method showed that grh was required in the R3 precursor for the correct acquisition of ommatidium polarity 20. (C) Visualization of mosaic crb[j1B5] mutant PRs, showing deformed homozygous mutant PRs. Crb is known to be required for rhabdomere morphogenesis.

 <img alt="Figure 6" fo:content-width="6in" src="/files/ftp_upload/50610/50610fig6.jpg" /> 
 Figure 6. Time-course analysis of mosaic fatp[k10307] mutant PRs with the Tomato/GFP-FLP/FRT method, over 14 days. A mosaic fatp[k10307] mutant retina is observed on day 1 (A, A'), day 4 (B, B'), day 8 (C, C') and day 14 (D, D') after hatching. The same group of mosaic PRs can be found at each time point, in the observed field (white rectangle, A, B, C, D). In this field of PRs (A', B', C', D'), homozygous mutant PRs are labeled in green, whereas heterozygous and homozygous wild-type PRs are labeled in yellow. From day 4 (B') onwards, the homozygous mutant PRs begin to disappear, indicating that the fatp[k10307] mutation induces a progressive degeneration of these PRs. <a href="https://www.jove.com/files/ftp_upload/50610/50610fig6large.jpg" target="_blank"> Click here to view larger figure</a>.

 <img alt="Figure 7" fo:content-width="6in" src="/files/ftp_upload/50610/50610fig7.jpg" /> 
 Figure 7. Rescue of fatp[k10307] mutant PRs with a mini-white-carrying fatp-expressing construct. The mosaic PRs are visualized with the Tomato/GFP-FLP/FRT method in 15-day-old flies. (A) Visualization of mosaic control PRs. (B) Visualization of mosaic fatp mutant PRs, showing the loss of mutant PRs. (C) Visualization of mosaic fatp mutant PRs in a fly reexpressing fatp in outer PR (rh1&gt;fatp). The mutant PRs are rescued. The presence of a mini-white gene and the associated red eye pigmentation in rh1&gt;fatp conditions does not alter Tomato or GFP fluorescence or clonal detection.

Watch this Scientific Journal Video about The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells at JoVE.com

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Tomato/GFP-FLP/FRT method involves visualizing mosaic photoreceptor cells in living Drosophila. It can be used to follow individual photoreceptor cell fates in the retina for days or weeks. This method is ideal for studies of retinal degeneration and neurodegenerative diseases or photoreceptor cell development.

The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

The Drosophila eye is widely used as a model  for studies of development and neuronal degeneration. With the powerful mitotic  recombination technique, ...

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18.0K vues.  Ecole Normale Supérieure de Lyon.  La méthode de tomate / GFP-FLP / FRT implique visualiser les cellules photoréceptrices de la mosaïque à vivre la drosophile. Il peut être utilisé pour suivre le destin des cellules individuelles des photorécepteurs dans la rétine pendant des jours ou des semaines. Ce procédé est idéal pour l'étude de la dégénérescence rétinienne et les maladies neurodégénératives ou le développement des cellules de photorécepteur. 

Vidéo: The Tomato/GFP-FLP/FRT Method for Live Imaging of Mosaic Adult Drosophila Photoreceptor Cells

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

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.

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

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

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.

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

Electron Spin Resonance Micro-imaging of Live Species for Oxygen Mapping

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment of live cells with micron-scale resolution1. Oxygen is one of the most important molecules in the cycle of life. It serves as the terminal electron acceptor of oxidative phosphorylation in the mitochondria and is used in the production of reactive oxygen species. Measurements of oxygen are important for the study of mitochondrial and metabolic functions, signaling pathways, effects of various stimuli, membrane permeability, and disease differentiation. Oxygen consumption is therefore an informative marker of cellular metabolism, which is broadly applicable to various biological systems from mitochondria to cells to whole organisms. Due to its importance, many methods have been developed for the measurements of oxygen in live systems. Current attempts to provide high-resolution oxygen imaging are based mainly on optical fluorescence and phosphorescence methods that fail to provide satisfactory results as they employ probes with high photo-toxicity and low oxygen sensitivity. ESR, which measures the signal from exogenous paramagnetic probes in the sample, is known to provide very accurate measurements of oxygen concentration. In a typical case, ESR measurements map the probe's lineshape broadening and/or relaxation-time shortening that are linked directly to the local oxygen concentration. (Oxygen is paramagnetic; therefore, when colliding with the exogenous paramagnetic probe, it shortness its relaxation times.) Traditionally, these types of experiments are carried out with low resolution, millimeter-scale ESR for small animals imaging. Here we show how ESR imaging can also be carried out in the micron-scale for the examination of small live samples. ESR micro-imaging is a relatively new methodology that enables the acquisition of spatially-resolved ESR signals with a resolution approaching 1 micron at room temperature2. The main aim of this protocol-paper is to show how this new method, along with newly developed oxygen-sensitive probes, can be applied to the mapping of oxygen levels in small live samples. A spatial resolution of ~30 x 30 x 100 &mu;m is demonstrated, with near-micromolar oxygen concentration sensitivity and sub-femtomole absolute oxygen sensitivity per voxel. The use of ESR micro-imaging for oxygen mapping near cells complements the currently available techniques based on micro-electrodes or fluorescence/phosphorescence. Furthermore, with the proper paramagnetic probe, it will also be readily applicable for intracellular oxygen micro-imaging, a capability which other methods find very difficult to achieve.

This protocol describes a method for micron-scale three-dimensional imaging of oxygen concentration in the immediate environment of live cells by electron spin resonance microscopy.

This protocol describes an electron spin resonance (ESR) micro-imaging method for three-dimensional mapping of oxygen levels in the immediate environment ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

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) ...