We thank an internal fund from the University of Oklahoma (to BZ) and a grant from NSF (IOS-1025556, to BZ and RAB) for supporting TRF, RAB, XP, AAO, MLW, CAS, and this research. An NIH grant (RO1 NS060878, to BZ) partially supported RAB. RAB acknowledges an NIH NRSA (T32 NS07292) training grant awarded to Brandeis University, and an NIH grant (RO1 GM21473) awarded to Jeffrey Hall at Brandeis, for supporting the early stages of this study. We thank Jeffrey Hall for his continuous encouragements for this project. We thank Dr. Kristin Scott for providing us with the tubP&gt;Gal80&gt; strain.

1. Drosophila strains 
<ol>
<li>Gal4 driver: 
GMR-Gal4, which expresses Gal4 in photoreceptor cells.</li>
<li>UAS lines 
UAS-MJD78Q (or UAS-polyQ), which causes neural degeneration in Gal4-expressing cells. 
UAS-GFP, which expresses GFP in Gal4-expressing cells.</li>
<li>FLP lines 
ey-FLP, which expresses FLP in photoreceptors and in subsets of brain regions. 
ET-FLPx2 lines, which are enhancer-trap lines containing two copies of FLP (FLP-IRES-FLP). We have generated &tilde; 1,000 ET-FLP lines (Bohm et al., 2010), whose expression patterns are yet to be characterized. Here, we describe the characterization of the expression pattern of select ET-FLPx2 in the larval and adult fly central nervous system (CNS).</li>
<li>FLP reporter 
yw, actinP&gt;CD2&gt;Gal4; UAS-GFP (Pignoni &amp; Zipursky, 1997), which reports the expression pattern of ET-FLPx2 through actinP&gt;Gal4; UAS-GFP upon FLP-mediated recombinational excision of &gt;CD2.</li>
<li>Two complementary Gal80-converting tools 
TubP&gt;Gal80&gt; (Gordon &amp; Scott, 2009), the flip-out construct, which constitutively expresses Gal80 in all tissues driven by a strong tubulin promoter. Upon FLP-mediated recombination, &gt;Gal80 is flipped out and Gal80 expression is terminated only in ET-FLPx2-expressing tissues. 
TubP&gt;stop&gt;Gal80 (Bohm et al., 2010), the flip-in construct, which does not express Gal80 under normal conditions. Gal80 expression is turned on by the tubulin promoter only in ET-FLPx2-expressing tissues in which FLP flips out &gt;stop and flips in Gal80.</li>
<li>Flies for testing FLP activity in the compound eye 
GMR-Gal4, UAS-polyQ; tubP&gt;stop&gt;Gal80 (Bohm et al., 2010), which has degenerate and depigmented eyes. In FLP-expressing photoreceptor cells, polyQ expression is turned off following Gal80 flip-in. 
GMR-Gal4, UAS-polyQ; tubP&gt;Gal80&gt; (Bohm et al., 2010), which has normal eyes. In FLP-expressing photoreceptor cells, polyQ expression is turned on following Gal80 flip-out.</li>
</ol>

2. Reagents
Phosphate-buffered saline (PBS, 1X), pH 7.4. 
4% paraformaldehyde (PFA), diluted in PBS from 16% PFA, EM grade (Cat#15710, Electronic Microscopy Sciences, Hatfield, PA) 
PBT, 1XPBS containing 0.1% Triton-X 100 (Sigma) 
Anti-Elav (mAb, 9F8A9; Developmental Studies Hybridoma Bank, University of Iowa), stains all neurons in Drosophila. 
Anti-Repo (mAb 8D12; Developmental Studies Hybridoma Bank, University of Iowa) stains glial cells in Drosophila. 
Alexa 594 chicken against mouse secondary antibody (Invitrogen)

3. Mapping the expression pattern of enhancer-trap FLPx2 lines (* critical steps)
<ol>
<li>Collect virgins females from the yw, actinP&gt;CD2&gt;Gal4; UAS-GFP line and cross to males of individual ET-FLPx2 lines. The former serves as a GFP reporter for tissue expression of ET-FLP.</li>
<li>*Dissect wandering 3rd instar F1 larvae, retain the CNS (i.e., the brain and the ventral nerve cord, VNC) and the body wall, fix for 1 hour in 4% PFA on ice, wash 3X with ice-cold PBS followed by 3X washes with PBT. 
Dissection
<ol>
<li>Remove wandering 3rd instar F1 larvae and place them into a clean Sylgard dish (35 mm x 10 mm petri dish filled with 2 grams of Sylgard - mixed according to manufacture instructions).
<ol>
	<li>Remove food debris with paintbrush</li></ol></li>
<li>Fill the dish with ice-cold 1XPBS and place on ice to anesthetize the larvae</li>
<li>Hold the mouth hooks of a larva with a pair of forceps and grab the cuticle near the extending anterior spiracle with another pair of forceps and peel the cuticle towards the posterior end. The CNS will be attached to the mouth hooks along with the salivary glands, gut, and imaginal discs.</li>
<li>Remove the excess tissue and imaginal discs surrounding the CNS.</li>
<li>Transfer the CNS into a PBS-filled 3 well pyrex dish.<ol>
		<li>A siliconized P200 pipette tip can be used to avoid drying the tissue.</li></ol></li> 
<li>Fix for 1 hour in 200 &mu;l of 4% PFA on ice.<ol>	
		<li>Ensure that the CNS is fully submerged in the PFA.<ol>
				<li>If the CNS floats, this typically suggests that the CNS is not properly cleaned from the surrounding tissue.</li></ol></li></ol></li> 
<li>Wash 3X with ice-cold PBS followed by 3X washes with PBT.</li>
</ol>
</li>
<li>*Dissect the CNS of adult F1 flies.
<ol>
<li>Anesthetize the flies with CO2 or place the flies contained inside an empty vial on ice for 10 minutes.</li>
<li>Place adult flies into a 95% EtOH-filled glass dish on ice for 30 sec.</li>
<li>Transfer the adult flies into a 1XPBS filled glass dish on ice.<ol>
		<li>Wash 3X with ice-cold 1X PBS to remove residual EtOH</li></ol></li> 
<li>Place a fly into a 35mm Sylgard dish filled with ice-cold 1X PBS</li>
<li>Place the ventral side facing up and insert a small insect pen (minutiens pins 0.1 mm) midway through the abdomen.<ol>
		<li>Using a pair of forceps hold the base of the legs and remove the legs with another pair of forceps.</li></ol></li>
<li>Remove the head cuticle<ol>
	<li>While grabbing the proboscis with one pair of forceps, place another underneath the eye and peel the cuticle towards the dorsal mid-line of the fly head.</li> 
	<li>Grab the other eye and remove the remaining cuticle on the other side.</li> 
	<li>Using sharp forceps and/or sharpened tungsten needle, remove tracheal tissue surrounding the brain<ol>
	<li>Improper cleaning of the trachea will result in strong auto-fluorescence, obstructing GFP expression, and allow the brain to float during fixing</li></ol></li></ol></li>
<li>VNC Dissection<ol>
		<li>Remove the thoracic cuticle<ol>
				<li>Starting at the posterior end of the thorax, place two forceps on each side of the 3rd preepisternum (i.e., grabbing the base of the hind legs).</li>
				<li>Gently separate the cuticle away from the ventral midline line, repeat this until you reach the base of the neck.</li> 
				<li>Gently spread the two halves of thoracic cuticle exposing the VNC</li></ol></li>
		<li>Next remove the abdomen from the thorax. While using a pair of forceps to hold the thorax at the second leg base, use another pair of forceps to grab the abdomen near the 1st abdominal sternite and pull the abdomen off.</li>
		<li>To remove the VNC from the surrounding thoracic tissue. Hold the humeral area with a pair of forceps and with another pair of forceps gentle tear away the majority of the thoracic carcass.</li> 
		<li>At this point there should be a ring of connective tissue surrounding the cervical connective (neck). Using two forceps tear the ring apart.</li> 
		<li>Remove any excess thoracic tissue surrounding the VNC</li></ol></li> 
<li>Transfer the brain and VNC to 9-well Pyrex dish containing 1X PBS on ice<ol>
		<li>A siliconized P200 pipette tip can be used to avoid drying the tissue</li></ol></li>
</ol>
</li>
<li>*Fix the adult CNS for 1 hour in 4% PFA on ice and wash 3X with ice-cold PBS and 3X with PBT.</li>
<li>Incubate the adult CNS preparation or the larval CNS and body wall at 4&deg;C overnight with antibodies to either neurons (anti-Elav, 1:10) or glia (anti-Repo. 1:10). Wash the preparation 5X with PBT.</li>
<li>Incubate with adult or larval preparation with Alexa 594 chicken against mouse secondary antibody (1:100) at room temperatures for 2 hrs, and wash the preparation with PBS.</li>
<li>Mount the larval or adult CNS on a slide. To preserve the natural shape of the CNS, use an O-shaped plastic ring (Clear Reinforcement Label, Avery Dennison, Office Products, Brea, CA) on the slide as a mini-bridge to suspend the cover slip.</li>
<li>Examine the preparation for GFP and Elav (or Repo) expression patterns under a fluorescent microscope.</li>
</ol>
4. The application of two complementary Gal80-converting tools in repression Gal4 in photoreceptors
<ol>
<li>Flipping Gal80 out via tubP&gt;Gal80&gt; (Figure 5A).<ol>
		<li>Collect males from the GMR-Gal4, UAS-polyQ; tubP&gt;Gal80&gt; line. Note that the compound eyes are normal in these flies.</li>
		<li>Mate with ey-FLP virgins.</li>
		<li>Collect the F1 flies and examine the eye phenotype.</li></ol></li>
<li>Flipping Gal80 in via tubP&gt;stop&gt;Gal80 (Figure 5B).<ol>
 		<li>Collect GMR-Gal4, UAS-polyQ; tubP&gt;stop&gt;Gal80 male flies and examine the eye phenotype. Note the degenerate and depigmented eyes in these flies compared to wild type strains (Canton-S)</li> 
	<li>Mate with ey-FLP virgins;</li>
	<li>Collect the F1 flies and examine the eye phenotype.</li></ol></li> 
<li>Select ET-FLPx2 lines with expression in photoreceptor cells
	</li> 
</ol>
Steps 4.1 and 4.2 can be repeated using ET-FLPx2 lines that express FLP in a subset of photoreceptor cells. Alternatively, either the GMR-Gal4, UAS-polyQ; tubP&gt;stop&gt;Gal80 flies or the GMR-Gal4, UAS-polyQ; tubP&gt;Gal80&gt; flies can be crossed to individual ET-FLPx2 lines to screen for lines that express FLP in photoreceptor cells. Valuable ET-FLPx2 lines are those that only express FLP in a subset or single photoreceptor cells (Figure 6).
5. Representative results
The expression pattern of a specific ET-FLPx2 line can be readily examined and documented in the F1 larvae or adult flies derived from the cross between yw, actinP&gt;CD2&gt;Gal4; UAS-GFP virgins and ET-FLPx2 males (Figure 1). Typically, three to five replicates of the F1 animals are dissected and examined to determine the reproducibility of the FLP expression pattern (Bohm et al., 2010). Figure 2 shows the expression pattern of FLP in ET-FLPx2 line 173A in the larval and adult CNS. The FLP pattern can be further analyzed using a neuronal or glial marker (Figure 3 &amp; 4). 
The results shown in Figure 7 illustrate the effectiveness of Gal80 flip-in and flip-out systems in intersectional repression of Gal4 following FLP-mediated recombination. The tubP&gt;stop&gt;Gal80 construct does not appear to have any leaky expression of Gal80 (based on the uniform degeneration and depigmentation of photoreceptors), and yet it is 100% effective in flipping Gal80 in with the help of eyFLP to repress GMR-Gal4 and restore the eye to normal appearances (Fig. 7A). On the contrary, tubP&gt;Gal80&gt; fully represses GMR-Gal4 prior to the introduction of eyFLP. Following Gal80 flip-out, GMR-Gal4 drives UAS-polyQ expression, leading to degenerate and depigmented eyes (Fig. 7B). In this example, we used a pan-photorecetor FLP to map the photoreceptor 'circuits' but did not examine the behavioral consequences. Using similar approaches with behaviorally relevant Gal4 lines, we mapped part of the CCAP-Gal4 circuit regulating wing expansion in adult flies (Bohm et al., 2010). We believe that the FINGR method will be valuable in aiding our understanding of the neuronal basis of behavior.
<img src="/files/ftp_upload/2649/2649fig1.jpg" alt="Figure 1"/> 
Figure 1. Examining the expression pattern of an ET-FLPx2 line. The FLP expression pattern (blue ovals) from an ET-FLPx2 line can be determined by crossing an ET-FLPx2 male (top left) with an actinP&gt;CD2&gt;Gal4; UAS-GFP virgin (top right). Ubiquitous ActinP&gt;Gal4 expression in the parental female is hindered by the insertion of the FRT (grey triangles) flanked CD2 sequence. In the F1 progeny, FLP will excise the CD2 sequence, permitting ActinP&gt;Gal4 to drive UAS-GFP expression.
<img src="/files/ftp_upload/2649/2649fig2.jpg" alt="Figure 2"/> 
Figure 2. Expression pattern of an ET-FLPx2 line in the larval and adult central nervous system. These images depict the CNS (brain and ventral nerve cord, VNC) dissected from the offspring produced from mating yw, actinP&gt;CD2&gt;Gal4; UAS-GFP parents to the ET-FLPx2 173A line. By recombining out the &gt;CD2&gt; cassette, FLP activates actinP&gt;GAL4 expression (the consequent expression of UAS-GFP). Hence, the GFP pattern reports FLP-expressing cells. Panel A shows the larval FLP expression pattern whereas panel B shows the adult expression pattern in line 173A.
<img src="/files/ftp_upload/2649/2649fig3.jpg" alt="Figure 3"/> 
Figure 3. Characterization of cell types which express flippase in ET-FLPx2 Line #688A. A, A composite image of a fixed F1 3rd instar larva stained with anti-REPO from a cross between ET-FLPx2 line #688A and a GFP Flippase reporting line. B-D, A high magnification (Zeiss Plan-Apochromat 63x oil) view taken from the posterior end of a VNC from a 3rd instar larvae.
<img src="/files/ftp_upload/2649/2649fig4.jpg" alt="Figure 4"/> 
Figure 4. Characterization of cell types within the mushroom body of ET-FLPx2 Line #150. A-C, A high-magnification image (63x oil immersion) of the mushroom body from a fixed F1 3rd instar larva stained with anti-REPO. 
<img src="/files/ftp_upload/2649/2649fig5.jpg" alt="Figure 5"/> 
Figure 5. Illustration of Gal80 Flip-in and Flip-out Strategies. A. In the flip-out strategy, tubulin promoter-driven Gal80 (TubP&gt;Gal80&gt;) is constitutively active, resulting in the repression of GMR-Gal4 and a wild-type eye phenotype (red eye). This repression is removed by the introduction of FLP (purple), which will flip Gal80 out (middle, red) and allow GMR-Gal4 to drive UAS-polyQ expression. This results in photoreceptor cell degeneration (illustrated by white dots in the eye). B. In the reciprocal of the previous strategy, tubulin promoter-driven Gal80 expression is interrupted by FRT (grey triangle) flanked "stop" codon. In the absence of Gal80, GMR-Gal4 binds to UAS-polyQ causing neuronal degeneration in photoreceptors (illustrated by white dots in the eye). By crossing the GMR-Gal4, UAS-polyQ; tubP&gt;stop&gt;GAL80 males with virgin females which express FLP in the photoreceptor cells, FLP (purple) will catalyze recombination at the FRT sites, excising the "stop" codon (middle, red), thus allowing for Gal80 expression and repression of GMR-Gal4. This reserves the neurodegeneration in photoreceptor cells (illustrated by the lack of white dots in the eye).
<img src="/files/ftp_upload/2649/2649fig6.jpg" alt="Figure 6"/> 
Figure 6. The application of the FINGR method in flipping Gal80 out using ET-FLPx2 Line #688A. F1 progeny from a cross between ET-FLPx2 line #688A virgin to GMR-Gal4, UAS-polyQ; tubP&gt;GAL80 male. The lack of pigment in the middle of the eye ommatidia represents FLP expression in line #688A. In this area Gal80 suppression of GMR-Gal4 was relieved allowing the Gal4 to drive UAS-polyQ.
<img src="/files/ftp_upload/2649/2649fig7.jpg" alt="Figure 7"/> 
Figure 7. The application of the FINGR method in flipping Gal80 in or out to intersect Gal4 in photoreceptors. A. The tubP&gt;stop&gt;Gal80 flip-in system restricts GAL4 expression in an FLP-dependent manner. In GMR-Gal4, UAS-polyQ; tubP&gt;stop&gt;GAL80 flies, photoreceptor cells are degenerate and depigmented (left fly). Following the introduction of eyFLP, Gal80 is expressed in photoreceptor cells, resulting in full repression of GMR-Gal4, which in turn turns off the expression polyQ and produces flies with normal eyes (right fly). B. The tubP&gt;Gal80&gt; flip-out system works in the opposite fashion as the tubP&gt;stop&gt;Gal80 flip-in system. TubP&gt;Gal80&gt; constitutively represses GAL4 and suppresses the expression of polyQ (left fly, with normal eyes). Following the introduction of eyFLP, Gal80 is flipped out from photoreceptor cells, relieving the suppression of GMR-Gal4 and turning on the expression polyQ. This leads to flies with degenerate and depigmented eyes (right fly).

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	<li>Bohm, R. A., Welch, W. P., Goodnight, L. K., Cox, L. W., Henry, L. G., Gunter, T. C., Bao, H., Zhang, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+genetic+mosaic+approach+for+neural+circuit+mapping+in+Drosophila.">A genetic mosaic approach for neural circuit mapping in Drosophila.</a> Proc Natl Acad Sci U S A. 107, 16378-16383 (2010).</li><li>Brand, A. H., Perrimon, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+gene+expression+as+a+means+of+altering+cell+fates+and+generating+dominant+phenotypes.">Targeted gene expression as a means of altering cell fates and generating dominant phenotypes.</a> Development. 118, 401-415 (1993).</li><li>Bruno, B., Holbro, N., Reichert, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polycomb+group+genes+are+required+for+neural+stem+cell+survival+in+postembryonic+neurogenesis+of+Drosophila.">Polycomb group genes are required for neural stem cell survival in postembryonic neurogenesis of Drosophila.</a> Development. 134, 1091-1099 (2007).</li><li>Duffy, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GAL4+system+in+Drosophila:+a+fly+geneticist's+Swiss+army+knife.">GAL4 system in Drosophila: a fly geneticist's Swiss army knife.</a> Genesis. , 34-341 (2002).</li><li>Gordon, M. D., Scott, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+control+in+a+Drosophila+taste+circuit.">Motor control in a Drosophila taste circuit.</a> Neuron. 61, 373-384 (2009).</li><li>Lee, T., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosaic+analysis+with+a+repressible+cell+marker+for+studies+of+gene+function+in+neuronal+morphogenesis.">Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis.</a> Neuron. 22, 451-461 (1999).</li><li>McGuire, S. E., Le, P. T., Osborn, A. J., Matsumoto, K., Davis, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiotemporal+rescue+of+memory+dysfunction+in+Drosophila.">Spatiotemporal rescue of memory dysfunction in Drosophila.</a> Science. 302, 1765-1768 (2003).</li><li>Osterwalder, T., Yoon, K. S., White, B. H., Keshishian, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+conditional+tissue+specific+transgene+expression+system+using+inducible+GAL4.">A conditional tissue specific transgene expression system using inducible GAL4.</a> Proc Natl Acad Sci USA. 98, 12596-12601 (2001).</li><li>Pignoni, F., Zipursky, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+Drosophila+eye+development+by+decapentaplegic.">Induction of Drosophila eye development by decapentaplegic.</a> Development. 124, 271-278 (1997).</li><li>Roman, G., Endo, K., Zong, L., Davis, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=P[Switch],+a+system+for+spatial+and+temporal+control+of+gene+expression+in+Drosophila+melanogaster.">P[Switch], a system for spatial and temporal control of gene expression in Drosophila melanogaster.</a> Proc Natl Acad Sci U S A. 98, 12602-12607 (2001).</li><li>Xu, T., Rubin, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+genetic+mosaics+in+developing+and+adult+Drosophila+tissues.">Analysis of genetic mosaics in developing and adult Drosophila tissues.</a> Development. 117, 1223-1237 (1993).</li></ol>

No conflicts of interest declared.

The Gal4/UAS system has been widely and successfully used by Drosophilists for various biological studies (Brand &amp; Perrimon, 1993; Duffy, 2002). To date, the fly community has generated thousands of promoter-driven and enhancer-trap Gal4 lines. A major strength of the FINGR method is that it is compatible with the Gal4/UAS system, making it possible to significantly expand the power of Gal4 in Drosophila studies. 
One important application of the FINGR method is to fine tune neural circuits underlying behavior through Gal4/Gal80 intersection. These behaviors may include, but are not limited to, learning and memory, courtship, sleep, and circadian rhythm. The FINGR method can also be used to map the critical neurons that are prone to neurodegeneration in modeling human neurodegenerative diseases such as Alzheimer's, amyotrophic lateral sclerosis (ALS), and Parkinson's diseases. Similarly, foci of neurons or neural circuits crucial for mediating cocaine, alcohol, and other substance addiction might be mapped using the FINGR method. It should be noted that conditional Gal4s (Osterwalder et al., 2001; Roman et al., 2001) could be used with the FINGR method in order to gain temporal control of neural circuits and behaviors. Modification of the Gal80-converting tools by using a temperature-sensitive Gal80 (Gal80ts, McGuire et al., 2003) can produce tubP&gt;stop&gt;Gal80ts or tubP&gt;Gal80ts&gt;, adding further temporal flexibilities in circuit manipulation. Beyond the nervous system, the FINGR method is equally applicable to restricting Gal4 expression in subsets of cells whether it is the wing or the leg. 
The ET-FLPx2 lines will be a valuable resource for all Drosophila biologists interested in FRT/FLP-based clonal analysis (Xu &amp; Rubin, 1993), including MARCM (Lee &amp; Luo, 1999). It is also expected that some ET-FLPx2 lines may replace heatshock-FLP for producing reproducible clones and for repeated morphological and behavioral analyses. Like enhancer-trap Gal4 lines, most ET-FLPx2 lines are unlikely to be expressed specifically in the cells of one's interest. Most lines will express FLP in the cells of interest AND in other cells or tissues as well. However, the lack of 'tissue specificity' of both Gal4 and ET-FLPx2 will not be a major concern for the FINGR method as long as their overlapping region is desirable for a particular experiment. FINGR is designed to turn two 'broad' systems into a refined one.
The methods illustrated in these protocols are rather standard and should be reproducible. The major cautionary steps that require one's attention are the selection and usage of the fixative. Fixatives other than PFA should be avoided because they will quench GFP signal. We have also noted that the dissection dish should not be used for fixation with PFA to avoid the possibility that FLP expression will not be reliably reported by GFP.

mapping and application of enhancer-trap flippase expression in larval and adult drosophila cns

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Watch this Scientific Journal Video about Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS at JoVE.com

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

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Neuroscience

16.4K vistas.  University of Oklahoma - Norman. Se describe un Flippase inducida intersectorial Gal80/Gal4 represión (FINGR) método, permitiendo al tejido-específica para determinar FLP GAL80 patrones de expresión. Siempre que sea Gal4 y la superposición de FLP, Gal4 expresión es activada (GAL80 un tirón hacia fuera) o desactivar (GAL80 volcó en). El método FINGR es versátil para el análisis clonal y la asignación de circuitos neuronales.

Video: Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific regions of developing embryos, including those of C. elegans, chick, Xenopus, zebrafish, and mouse 1. In zebrafish, in vivo electroporation has been shown to have excellent spatial and temporal resolution for the delivery of these reagents 2-7. The temporal resolution of this method is important because it allows for incorporation of these reagents at specific stages in development. Furthermore, because expression from electroporated vectors occurs within 6 hours 7, this method is more timely than transgenic approaches. While the spatial resolution can be extremely precise when targeting a single cell 2, 6, it is often preferable to incorporate reagents into a specific cell population within a tissue or structure. When targeting multiple cells, in vivo electroporation is efficient for delivery to a specific region of the embryo; however, particularly within the developing nervous system, it is difficult to target specific cell types solely through spatially discrete electroporation. Alternatively, enhancer trap transgenic lines offer excellent cell type-specific expression of transgenes 8. Here we describe an approach that combines transgenic Gal4-based enhancer trap lines 8 with spatially discrete in vivo electroporation 7, 9 to specifically target developing neurons of the zebrafish olfactory bulb. The Et(zic4:Gal4TA4,UAS:mCherry)hzm5 (formerly GA80_9) enhancer trap line previously described 8, displays targeted transgenic expression of mCherry mediated by a zebrafish optimized Gal4 (KalTA4) transcriptional activator in multiple regions of the developing brain including hindbrain, cerebellum, forebrain, and the olfactory bulb. To target GFP expression specifically to the olfactory bulb, a plasmid with the coding sequence of GFP under control of multiple Gal4 binding sites (UAS) was electroporated into the anterior end of the forebrain at 24-28 hours post-fertilization (hpf). Although this method incorporates plasmid DNA into multiple regions of the forebrain, GFP expression is only induced in cells transgenically expressing the KalTA4 transcription factor. Thus, by using the GA080_9 transgenic line, this approach led to GFP expression exclusively in the developing olfactory bulb. GFP expressing cells targeted through this approach showed typical axonal projections, as previously described for mitral cells of the olfactory bulb 10. This method could also be used for targeted delivery of other reagents including short-hairpin RNA interference expression plasmids, which would provide a method for spatially and temporally discrete loss-of-function analysis.

Targeting Olfactory Bulb Neurons Using Combined In Vivo Electroporation and Gal4-Based Enhancer Trap Zebrafish Lines

The temporal and spatial resolution of genetic manipulations determines the spectrum of biological phenomena that they can perturb. Here we use temporally and spatially discrete in vivo electroporation, combined with transgenic lines of zebrafish, to induce expression of a GFP transgene specifically in neurons of the developing olfactory bulb.

In vivo electroporation is a powerful method for delivering DNA expression plasmids, RNAi reagents, and morpholino anti-sense oligonucleotides to specific ...

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

Drosophila Adult Olfactory Shock Learning

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the elucidation of the molecular mechanisms involved in cognitive diseases1-7. Learning and memory can be assayed in larvae to study the effect of neurodevelopmental genes8-10 and in flies to measure the contribution of adult plasticity genes1-7. Furthermore, the short lifespan of Drosophila facilitates the analysis of genes mediating age-related memory impairment5,11-13. The availability of many inducible promoters that subdivide the Drosophila nervous system makes it possible to determine when and where a gene of interest is required for normal memory as well as relay of different aspects of the reinforcement signal3,4,14,16.
Studying memory in adult Drosophila allows for a detailed analysis of the behavior and circuitry involved and a measurement of long-term memory15-17. The length of the adult stage accommodates longer-term genetic, behavioral, dietary and pharmacological manipulations of memory, in addition to determining the effect of aging and neurodegenerative disease on memory3-6,11-13,15-21.
Classical conditioning is induced by the simultaneous presentation of a neutral odor cue (conditioned stimulus, CS+) and a reinforcement stimulus, e.g., an electric shock or sucrose, (unconditioned stimulus, US), that become associated with one another by the animal1,16. A second conditioned stimulus (CS-) is subsequently presented without the US. During the testing phase, Drosophila are simultaneously presented with CS+ and CS- odors. After the Drosophila are provided time to choose between the odors, the distribution of the animals is recorded. This procedure allows associative aversive or appetitive conditioning to be reliably measured without a bias introduced by the innate preference for either of the conditioned stimuli. Various control experiments are also performed to test whether all genotypes respond normally to odor and reinforcement alone.

Drosophila Adult Olfactory Shock Learning

The method to measure adult Drosophila associative memory is described. The assay is based on the ability of the fly to associate an odor presented with a negative reinforcer (electric shock) and then recall this information at a later time, allowing memory to be measured.

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

Functional Imaging of Auditory Cortex in Adult Cats using High-field fMRI

Current knowledge of sensory processing in the mammalian auditory system is mainly derived from electrophysiological studies in a variety of animal models, including monkeys, ferrets, bats, rodents, and cats. In order to draw suitable parallels between human and animal models of auditory function, it is important to establish a bridge between human functional imaging studies and animal electrophysiological studies. Functional magnetic resonance imaging (fMRI) is an established, minimally invasive method of measuring broad patterns of hemodynamic activity across different regions of the cerebral cortex. This technique is widely used to probe sensory function in the human brain, is a useful tool in linking studies of auditory processing in both humans and animals and has been successfully used to investigate auditory function in monkeys and rodents. The following protocol describes an experimental procedure for investigating auditory function in anesthetized adult cats by measuring stimulus-evoked hemodynamic changes in auditory cortex using fMRI. This method facilitates comparison of the hemodynamic responses across different models of auditory function thus leading to a better understanding of species-independent features of the mammalian auditory cortex.

Functional studies of the auditory system in mammals have traditionally been conducted using spatially-focused techniques such as electrophysiological recordings. The following protocol describes a method of visualizing large-scale patterns of evoked hemodynamic activity in the cat auditory cortex using functional magnetic resonance imaging.

Current knowledge of sensory processing in the mammalian auditory system is mainly derived from electrophysiological studies in a variety of animal ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of the nervous system often produce viable larvae and adults that have locomotion defective phenotypes that are difficult to adequately describe with text or completely represent with a single photographic image. Current modes of scientific publishing, however, support the submission of digital video media as supplemental material to accompany a manuscript. Here we describe a simple and widely accessible microscopy technique for acquiring high-quality digital video of both Drosophila larval and adult phenotypes from a lateral perspective. Video of larval and adult locomotion from a side-view is advantageous because it allows the observation and analysis of subtle distinctions and variations in aberrant locomotive behaviors. We have successfully used the technique to visualize and quantify aberrant crawling behaviors in third instar larvae, in addition to adult mutant phenotypes and behaviors including grooming.

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Here we describe a simple and widely accessible microscopy technique to acquire high-quality digital video of Drosophila adult and larval mutant phenotypes from a lateral perspective.

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...