We thank Sean B. Carroll and Thomas Werner for providing fly stocks, Naoyuki Fuse for equipment, Byung Seok Jin for his assistance in filming, Kiyokazu Agata for mentoring and Elizabeth Nakajima for English editing. This work was supported by KAKENHI 17K19427 and Takeda Science Foundation.

1. Fly stock
<ol>
	<li>Use Drosophila guttifera for all of the following protocols.</li>
	<li>Use plastic vials (diameter 25 mm x height 96 mm) and cellulose plugs (diameter 23 mm x height 26 mm) for stock maintenance. Use a standard cornmeal/sugar/yeast/agar food and follow a publication described three other alternative recipes for this species2. 
	NOTE: D. guttifera (stock number 15130-1971.10) is provided by the Drosophila Species Stock Center at the University of California, San Diego. Although D. guttifera belongs to the immigrans-tripunctata radiation, which is distantly related to D. melanogaster within the genus23, it has many biological properties in common with D. melanogaster. Accordingly, this protocol can be applied for many Drosophila species, although some species require specific food and/or technical tips to maintain them2.</li>
</ol>
2. Observation of pupa and definition of pupal stages
NOTE: The pupa for observation is taken from the fly stock maintained with a 12:12 h light/dark cycle at 25 &#176;C. Bainbridge and Bownes17 described a low risk of moving D. melanogaster pupae from the original place of pupation onto a piece of moistened tissue paper (97% survival of 946 moved pupae). D. guttifera pupae can be prepared by essentially the same method.
<ol>
	<li>Place healthy adults of Drosophila on fresh food (standard cornmeal/sugar/yeast/agar food) in a plastic vial (diameter 25 mm x height 96 mm) and let them lay eggs. Wait 7 days to obtain late 3rd instars.</li>
	<li>Place 1 mL of standard cornmeal/sugar/yeast/agar food into each 1.5 mL microtube. Make 3 pinholes with 2 mm spacing in the lids of microtubes by penetrating with a push-pin to allow breathing.</li>
	<li>Centrifuge the microtubes with food for 7 s in a mini microcentrifuge (860 x g).</li>
	<li>Invert and tap vials to remove all adults from the vial.</li>
	<li>Pour 5 - 10 mL of ddH2O (or reverse osmosis water) into the vial.</li>
	<li>Pour out larvae with water into a plastic Petri dish (diameter 90 mm x height 15 mm). Identify late 3rd instars by their large body size (3 - 4 mm in length).</li>
	<li>Gently move late 3rd instar larvae with forceps into the microtubes with food (10 larvae / microtube). Incubate them overnight at 25 &#176;C.</li>
	<li>Move newly formed pupae onto a piece of tissue paper that has been moistened by ddH2O and placed in a plastic Petri dish (diameter 35 mm x height 10 mm).</li>
	<li>Place the Petri dish in a moist chamber (containing 10 mL of ddH2O in the bottom), and wait until the pupae develop to the desired stage.</li>
	<li>Move pupae onto a moistened piece of tissue paper in a plastic Petri dish (diameter 60 mm x height 15 mm). 
	NOTE: The definition of stages can mostly be made based on the stages of D. melanogaster17. Typically, the pupal period of Drosophila can be categorized into P1 - P15(ii), although some modifications of stage definition would be required depending on the species used. If possible, removing the puparium enables precise and detailed observation. See details below (Step 3).</li>
	<li>Observe pupae under a stereo microscope. Take photographs using a digital camera attached to the stereo microscope.</li>
</ol>
3. Removing puparium
NOTE: Pupae of Drosophila are covered by a structure called the puparium. An insect of Muscomorpha (flies) does not shed its larval cuticle at pupation; instead, it hardens the cuticle after apolysis, and uses it as a protective cover of the pupa, the puparium24. A pupa residing inside a puparium has a true pupal cuticle, which is very soft and fragile. Before apolysis takes place around P4(ii), epithelia and puparium are attached together, and therefore removing the puparium without damage is very difficult. After P5, removing the puparium is laborious, but useful for morphological observation and definition of pupal stages. The process is carried out as follows.
<ol>
	<li>Affix a piece of double-sided tape on a piece of paper towel.</li>
	<li>Place a pupa on the double-sided tape ventral side up (Figure 1A).</li>
	<li>Locate the space between the anterior side of the puparium and the internal pupa. Grasp and remove the puparium around this gap using forceps, and expose the anterior side of the head of the pupa.</li>
	<li>Insert the tip of a forceps by moving it parallel to the anterior-posterior axis. Lift the tip of the forceps to locally break the puparium. Repeat this action until the breakage reaches the posterior part of the puparium. Ensure that a gap is also formed between the puparium and pupal legs, and break the ventral side of the puparium and minimize the damage to the internal pupa (Figure 1B).</li>
	<li>After breaking the puparium as much as possible, take out the pupa using a fine paintbrush (#5/0) (Figure 1C).</li>
	<li>Place the pupa on a piece of tissue paper that has been moistened with ddH2O and placed in a plastic Petri dish (diameter 60 mm x height 15 mm). Take photographs as soon as possible because the exposed pupa is vulnerable and easily becomes dry. 
	NOTE: Pupae without a puparium are not suitable for measuring durations of pupal periods (Step 4), because stress (such as desiccation) and physical damage might interfere with the normal development.</li>
</ol>
4. Measuring durations of pupal stages
<ol>
	<li>Prepare pupae for measuring durations of pupal stages as described in Step 2. Collect pupae of 1 - 2, 2 - 3, and 3 - 4 days after pupal formation (20 pupae each). Give individual identification numbers (1 - 60) to pupae for identification. Continue collecting newly formed pupae during the following steps, to obtain 20 more young pupae. Give individual identification numbers (61 - 80) to the newly formed pupae. Place a pupa on a piece of ddH2O-moistened tissue paper in a well (3.9 cm2) of a 12-well cell culture plate (1 pupa/well, 12 pupae/plate). Fill the inter-well space of plates with ddH2O to maintain humidity, put the lids on, and place the plates in 25 &#176;C, constant light (24:0 h light/dark) conditions. 
	NOTE: Pupae without a puparium are not suitable for measuring durations of pupal periods. Please do not remove the puparium.</li>
	<li>Observe morphological features including body color, bristles, Malpighian tubules and yellow body of all pupae once every 30 min, and record observed stages (P1 - P15(ii), based on the references17,19) in a tally sheet.</li>
	<li>Continue recording over four straight days (96 h) by a rotating shift of three (or more) persons.</li>
	<li>Count numbers of records of each particular stage, and average them (average count of observations / pupae). Then multiply them by 0.5 (h), resulting in the estimated lengths of stages (h).</li>
</ol>
5. Measurement of intensity of black spots on a wing
NOTE: The intensity of black spots on a pupal or adult wing can be quantified by measuring optical density (OD). A glass filter with known ODs (stepped density filter) is used for calibration25, so that one can calculate the OD of a particular area from a digital image of a wing. The OD in a spot and the OD outside of the spot are measured, and the latter is subtracted from the former to obtain the intensity of the spot (&#916;OD). Here, we describe the method of dissection, measurement and calculation of &#916;OD. This procedure can be done after Step 2, independent from Step 3 and Step 4. Once one has performed Step 2 and understands all pupal stages, one can directly start or repeat Step 5.
<ol>
	<li>Image preparation
	<ol>
		<li>Prepare a new pupa of a focal stage as described in Step 2. Remove the anterior part of a puparium with forceps. Take out the pupa using forceps and place it into phosphate buffered saline (PBS, Table 2) in a plastic Petri dish (diameter 35 mm x height 10 mm).</li>
		<li>Cut the basal joint of a wing (basal joint is the narrow proximal part of the wing). As the wing is folded, place it into a plastic Petri dish (diameter 35 mm x height 10 mm) filled with ddH2O to extend it by osmotic pressure (the wing unfolds by itself).</li>
		<li>Collect newly eclosed adults once every 10 min from a stock vial. Anesthetize a fly with CO2 using a CO2 anesthetizing pad, confirm anesthetization by immobility and cut the basal joint of a wing.</li>
		<li>Place 10 &#181;L of PBS on a glass slide, place the wing there, and cover with a cover slip (18 mm x 18 mm).</li>
		<li>Turn on the light of the stereo microscope. Set the light to be at maximum level. Set the objective lens 11.5X. Set the diaphragm to be the most open state. Turn on the camera. Set the camera to be (ISO: 100, mode SHQ 3136 x 2352 pixels, shutter speed: 1/20 s). Focus on the sample by moving the focus knob of the microscope.</li>
		<li>Push the shutter button of the remote-control unit to take an image. Take 3 images per wing, each of which must be centered on a campaniform sensillum, longitudinal vein spot or posterior crossvein, positioning the distal part of the wing on the left side and the anterior part of the wing on the upper side.</li>
	</ol>
	</li>
	<li>Calibration
	<ol>
		<li>Take images of 9 parts of a stepped density filter using the same camera settings used to obtain the wing image.</li>
		<li>Initiate ImageJ software (https://imagej.nih.gov/ij/)22.</li>
		<li>Click File | Open | and select one of the images of the stepped density filter.</li>
		<li>Click Image | Type | 8-bit | to convert the image to an 8-bit image.</li>
		<li>Click Edit | Selection | Specify | and check Oval and Centered column. Write 100 (pixels) in Width column, 100 (pixels) in Height column, 1568 in X coordinate column and 1176 in Y coordinate column. Click OK.</li>
		<li>Click Analyze | Measure|. The &#34;mean grey value&#34; of selected areas are measured.</li>
		<li>Repeat 5.2.3. to 5.2.6. for the 8 remaining images.</li>
		<li>Click Analyze | Calibrate and select Rodbard25 in Function column and write the following number in the right column in the middle (0.04, 0.336, 0.632, 0.928, 1.224, 1.52, 1.816, 2.112, 2.408; these numbers depend on the densities of the stepped density filter).</li>
		<li>Check Global calibration column and click OK. 
		NOTE: By performing this procedure, &#34;mean grey value&#34; is converted to &#34;optical density (OD)&#34; using the Rodbard function. After this step, optical density can be calculated for a particular selected area in ImageJ software.</li>
	</ol>
	</li>
	<li>Choosing area of measurements 
	NOTE: Spots are typically associated with landmarks, such as campaniform sensilla, longitudinal vein tips, and crossveins. These and other landmarks on a wing can be used to choose the region of measurements. Here, an example in D. guttifera (Figure 2) is described.
	<ol>
		<li>Definition of Point A, campaniform sensillum spot.
		<ol>
			<li>Open an image in which a campaniform sensillum spot is at the center of the image. Click Image | Type | 8-bit | to convert the image to an 8-bit image.</li>
			<li>Click Rectangle in Area Selection Tools and draw a rectangle. Set the upper left vertex of the rectangle so that it is attached to the posterior line of the third longitudinal vein and more distal from a campaniform sensillum spot. Set the right side of the rectangle so that it is located to the right of the campaniform sensillum spot.</li>
			<li>Click Edit | Selection | Add to Manager |. Check Show All column.</li>
			<li>Click Angle tool in Line Selection Tools. Draw the first line on the posterior line of third longitudinal vein. Set the left endpoints of the line on the vertex of the rectangle drawn in Step 1. Draw the second line on the upper side of the rectangle. Press the &#34;m&#34; key to measure the angle between the two lines drawn in this step. Click the window of the image on the screen of the computer.</li>
			<li>Click Edit | Selection | Add to Manager |.</li>
			<li>Click Rectangle in Area Selection Tools and draw a rectangle of approximately 1/9 the size of the image window. Click Edit | Selection | Rotate |. Write the minus degrees of the angle measured in Step 5.3.1.4. in Angle column and click OK to rotate the rectangle drawn in this step.</li>
			<li>Move the rectangle drawn in Step 5.3.1.6. by using the arrow keys. Set the endpoint of the posterior line of the second longitudinal vein on the left side of the rectangle and the lower side of the rectangle attached to the posterior line of the third longitudinal vein. Confirm that a perpendicular line from the end point of the second longitudinal vein to the posterior line of the third longitudinal vein is drawn in this procedure. Define the foot of the perpendicular line as Point A (Figure 2A).</li>
			<li>Record the x coordinate and the y coordinate of Point A, indicated below Area Selection Tools when placing the cursor on Point A.</li>
		</ol>
		</li>
		<li>Definition of Point B, longitudinal vein tip spot
		<ol>
			<li>Open an image in which a longitudinal vein tip spot is at the center of the image. Click Image | Type | 8-bit | to convert the image to an 8-bit image.</li>
			<li>Repeat the same procedure described in Step 5.3.1. (Definition of Point A, campaniform sensillum spot) to find Point A in the image.</li>
			<li>Click Edit | Selection | Add to Manager |.</li>
			<li>Click Rectangle in Area Selection Tools and draw a rectangle. Set the upper left vertex of the rectangle at the end point of the posterior line of the third longitudinal vein.</li>
			<li>Click Edit | Selection | Add to Manager |.</li>
			<li>Click Angle tool in Line Selection Tools. Draw the first line to connect Point A and the end point of the posterior line of the third longitudinal vein. Define this line as Line A. Draw the second line on the upper side of the rectangle drawn in Step 5.3.2.4. Press &#34;m&#34; key to measure the angle between the two lines.</li>
			<li>Click Edit | Selection | Add to Manager |.</li>
			<li>Click Rectangle in Area Selection Tools and draw a rectangle of approximately 1/9 the size of the image window. Click Edit | Selection | Rotate |. Write the minus degrees of the angle measured in Step 5.3.2.6. in Angle column and click OK to rotate the rectangle drawn in this step. Move the rectangle by using the arrow keys. Set the upper side of the rectangle so that it is attached to Line A and the end point of the anterior line of the fourth longitudinal vein so that it is on the left side of the rectangle.</li>
			<li>Click Edit | Selection | Add to Manager |.</li>
			<li>Click Rectangle in Area Selection Tools and draw a rectangle of approximately 1/9 the size of the image window. Click Edit | Selection | Rotate |. Write the minus degrees of the angle measured in Step 6 in Angle column and click OK to rotate the rectangle drawn in this step. Move the rectangle by using the arrow keys. Set the lower left vertex of the rectangle so that it is at the upper left vertex of the rectangle drawn in Step 5.3.2.8., resulting in obtaining the perpendicular line from the end point of the anterior line of the fourth longitudinal vein to Line A. Define the intersection point of the perpendicular line and the posterior line of the third longitudinal vein as Point B (Figure 2B).</li>
			<li>Record the x coordinate and the y coordinate of Point B, indicated below Area Selection Tools when placing the cursor on Point B.</li>
		</ol>
		</li>
		<li>Definition of Point C, posterior crossvein spot
		<ol>
			<li>Open an image in which a posterior crossvein spot is at the center of the image. Click Image | Type | 8-bit | to convert the image to an 8-bit image.</li>
			<li>Define Point C as the posterior-most point of the anterior line of the fourth longitudinal vein in the intersection area of the posterior crossvein and the fourth longitudinal vein (Figure 2C).</li>
			<li>Record the x coordinate and the y coordinate of Point C, indicated below Area Selection Tools when placing the cursor on Point C.</li>
		</ol>
		</li>
		<li>Definition of Point D, control area
		<ol>
			<li>Open an image in which a campaniform sensillum spot is at the center of the image. Click Image | Type | 8-bit | to convert the image to an 8-bit image.</li>
			<li>Click Straight in Line Selection Tools and draw a line connecting the end point of the anterior line of the second longitudinal vein and the end point of the posterior line of the fourth longitudinal vein. Define Point D as the crossing point of this line and the posterior line of the third longitudinal vein (Figure 2D).</li>
			<li>Record the x coordinate and the y coordinate of Point D, indicated below Area Selection Tools when placing the cursor on Point D.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Measurements
	<ol>
		<li>Open one of the images of wing spots (for measurement of Point A, open the image with Point A in the center). Click Image | Type | 8-bit | to convert the image to an 8-bit image.</li>
		<li>Click Rectangle in Area Selection Tools and draw a rectangle of approximately 1/9 size of the image window.</li>
		<li>Click Edit | Selection | Specify |. Check Oval and Centered column. Write 100 (pixels) in Width and Height columns, write the x coordinates of Point A (or Point B or Point C) and D in X coordinate column and write the y coordinates of Point A (or Point B or Point C) and D in Y coordinate column. Click OK.</li>
		<li>Click Analyze | Measure |. If the calibration described in 5.2 has already been finished, ODs are indicated in Mean column.</li>
		<li>Calculate &#916;ODs by subtracting OD of Point D from ODs of Points A, B, and C. 
		NOTE: Point D is in a transparent part of the wing and does not include pigmentation, and therefore is suitable for a background control.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Carson, H. L., Hardy, D. E., Spieth, H. T., Stone, W. S., Hecht, M. K., Steere, W. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+evolutionary+biology+of+the+Hawaiian+Drosophilidae.">The evolutionary biology of the Hawaiian Drosophilidae.</a> Essays in evolution and genetics in honor of Theodosius Dobzhansky. , 437-543 (1970).</li><li>Markow, T. A., O'Grady, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophila: a guide to species identification and use. , (2006).</li><li>Patterson, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Drosophilidae of the southwest. 4313, 7-216 (1943).</li><li>Setoguchi, S., Takamori, H., Aotsuka, T., Sese, J., Ishikawa, Y., Matsuo, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sexual+dimorphism+and+courtship+behavior+in+Drosophila+prolongata.">Sexual dimorphism and courtship behavior in Drosophila prolongata.</a> J Ethol. 32 (2), 91-102 (2014).</li><li>Werner, T., Jaenike, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophilids of the Midwest and Northeast. , (2017).</li><li>Arnoult, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Emergence+and+diversification+of+fly+pigmentation+through+evolution+of+a+gene+regulatory+module.">Emergence and diversification of fly pigmentation through evolution of a gene regulatory module.</a> Science. 339 (6126), 1423-1426 (2013).</li><li>Camino, E. M., Butts, J. C., Ordway, A., Vellky, J. E., Rebeiz, M., Williams, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+evolutionary+origination+and+diversification+of+a+dimorphic+gene+regulatory+network+through+parallel+innovations+in+cis+and+trans.">The evolutionary origination and diversification of a dimorphic gene regulatory network through parallel innovations in cis and trans.</a> PLoS Genet. 11 (4), e1005136 (2015).</li><li>Gompel, N., Prud'homme, B., Wittkopp, P. J., Kassner, V. A., Carroll, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chance+caught+on+the+wing:+cis-regulatory+evolution+and+the+origin+of+pigment+patterns+in+Drosophila.">Chance caught on the wing: cis-regulatory evolution and the origin of pigment patterns in Drosophila.</a> Nature. 433 (7025), 481-487 (2005).</li><li>Glassford, W. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Co-option+of+an+ancestral+Hox-regulated+network+underlies+a+recently+evolved+morphological+novelty.">Co-option of an ancestral Hox-regulated network underlies a recently evolved morphological novelty.</a> Dev. Cell. 34 (5), 520-531 (2015).</li><li>Koshikawa, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enhancer+modularity+and+the+evolution+of+new+traits.">Enhancer modularity and the evolution of new traits.</a> Fly. 9 (4), 155-159 (2015).</li><li>Koshikawa, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gain+of+cis-regulatory+activities+underlies+novel+domains+of+wingless+gene+expression+in+Drosophila.">Gain of cis-regulatory activities underlies novel domains of wingless gene expression in Drosophila.</a> Proc Natl Acad Sci U S A. 112 (24), 7524-7529 (2015).</li><li>McGregor, A. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+evolution+through+multiple+cis-regulatory+mutations+at+a+single+gene.">Morphological evolution through multiple cis-regulatory mutations at a single gene.</a> Nature. 448 (7153), 587-590 (2007).</li><li>Tanaka, K., Barmina, O., Kopp, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+developmental+mechanisms+underlie+the+evolutionary+diversification+of+Drosophila+sex+combs.">Distinct developmental mechanisms underlie the evolutionary diversification of Drosophila sex combs.</a> Proc Natl Acad Sci U S A. 106 (12), 4764-4769 (2009).</li><li>Werner, T., Koshikawa, S., Williams, T. M., Carroll, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+a+novel+wing+colour+pattern+by+the+Wingless+morphogen.">Generation of a novel wing colour pattern by the Wingless morphogen.</a> Nature. 464 (7292), 1143-1148 (2010).</li><li>Wittkopp, P. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intraspecific+polymorphism+to+interspecific+divergence:+genetics+of+pigmentation+in+Drosophila.">Intraspecific polymorphism to interspecific divergence: genetics of pigmentation in Drosophila.</a> Science. 326 (5952), 540-544 (2009).</li><li>Lawrence, P. A., Morata, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compartments+in+the+wing+of+Drosophila:+a+study+of+the+engrailed+gene.">Compartments in the wing of Drosophila: a study of the engrailed gene.</a> Dev Biol. 50 (2), 321-337 (1976).</li><li>Bainbridge, S. P., Bownes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Staging+the+metamorphosis+of+Drosophila+melanogaster.">Staging the metamorphosis of Drosophila melanogaster.</a> J. Embryol. Exp. Morphol. 66, 57-80 (1981).</li><li>Ashburner, M., Golic, K. G., Hawley, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophila: A Laboratory Handbook. , (2005).</li><li>Fukutomi, Y., Matsumoto, K., Agata, K., Funayama, N., Koshikawa, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pupal+development+and+pigmentation+process+of+a+polka-dotted+fruit+fly,+Drosophila+guttifera+(Insecta,+Diptera).">Pupal development and pigmentation process of a polka-dotted fruit fly, Drosophila guttifera (Insecta, Diptera).</a> Dev Genes Evol. 227 (3), 171-180 (2017).</li><li>Koshikawa, S., Fukutomi, Y., Matsumoto, K., Sekimura, T., Nijhout, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+guttifera+as+a+model+system+for+unraveling+color+pattern+formation.">Drosophila guttifera as a model system for unraveling color pattern formation.</a> Diversity and evolution of butterfly wing patterns: an integrative approach. , (2017).</li><li>True, J. R., Edwards, K. A., Yamamoto, D., Carroll, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+wing+melanin+patterns+form+by+vein-dependent+elaboration+of+enzymatic+prepatterns.">Drosophila wing melanin patterns form by vein-dependent elaboration of enzymatic prepatterns.</a> Curr Biol. 9 (23), 1382-1391 (1999).</li><li>Schneider, C. A., Rasband, W. S., Eliceiri, K. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NIH+Image+to+ImageJ:+25+years+of+image+analysis.">NIH Image to ImageJ: 25 years of image analysis.</a> Nat Methods. 9 (7), 671-675 (2012).</li><li>Izumitani, H. F., Kusaka, Y., Koshikawa, S., Toda, M. J., Katoh, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phylogeography+of+the+Subgenus+Drosophila+(Diptera:+Drosophilidae):+evolutionary+history+of+faunal+divergence+between+the+Old+and+the+New+Worlds.">Phylogeography of the Subgenus Drosophila (Diptera: Drosophilidae): evolutionary history of faunal divergence between the Old and the New Worlds.</a> PLoS ONE. 11 (7), e0160051 (2016).</li><li>Resh, V. H., Card&#233;, R. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Encyclopedia of Insects. , (2009).</li><li>DeLean, A., Munson, P. J., Rodbard, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+analysis+of+families+of+sigmoidal+curves:+application+to+bioassay,+radioligand+assay,+and+physiological+dose-response+curves.">Simultaneous analysis of families of sigmoidal curves: application to bioassay, radioligand assay, and physiological dose-response curves.</a> Am J Physiol. 235 (2), E97-E102 (1978).</li><li>Robertson, C. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+metamorphosis+of+Drosophila+melanogaster,+including+an+accurately+timed+account+of+the+principal+morphological+changes.">The metamorphosis of Drosophila melanogaster, including an accurately timed account of the principal morphological changes.</a> J Morphol. 59 (2), 351-399 (1936).</li><li>McKinney, M. L., McNamara, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Heterochrony: the evolution of ontogeny. , (1991).</li><li>Hardie, D. C., Gregory, T. R., Hebert, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+pixels+to+picograms:+a+beginners'+guide+to+genome+quantification+by+Feulgen+image+analysis+densitometry.">From pixels to picograms: a beginners' guide to genome quantification by Feulgen image analysis densitometry.</a> J Histochem Cytochem. 50 (6), 735-749 (2002).</li><li>Koshikawa, S., Miyazaki, S., Cornette, R., Matsumoto, T., Miura, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome+size+of+termites+(Insecta,+Dictyoptera,+Isoptera)+and+wood+roaches+(Insecta,+Dictyoptera,+Cryptocercidae).">Genome size of termites (Insecta, Dictyoptera, Isoptera) and wood roaches (Insecta, Dictyoptera, Cryptocercidae).</a> Naturwissenschaften. 95 (9), 859-867 (2008).</li></ol>

The authors have no conflict of interest.

We describe here the protocols for definition of pupal stages, removing the puparium for detailed observation, measuring durations of pupal stages, and measurement of intensity of black spots on a wing in D. guttifera. These protocols can be applied for many Drosophila and related fly species, especially species with wing pigmentation.
In-depth observation and description of more detailed developmental events would enable further subdivision of stages. In many cases, a develo...

Some of the morphological traits of Drosophila are diversified among species1,2,3,4,5. We can approach the question of how morphological diversity arises by comparing the mechanisms of generation of these morphologies. Examples of such morphologies are larval trichomes, adult sex combs, external genital apparatus, abdominal pigmentation, and wing pigmentation6,7,8,9,10,11,12,13,14,15. To study morphological differences among adults, observation and analysis of the pupal stages are important, because the fate of adult traits is determined in the late larval stages and subsequent morphogenesis proceeds during the pupal period.
In developmental biology studies of Drosophila melanogaster, &#34;hours APF&#34; (hours after pupal formation) is the common method to indicate a pupal stage16. This system employs absolute time after pupal formation and is very convenient for routine experiments. However, developmental speed may differ among pupae, and may be affected by slight genetic, epigenetic or microenvironmental differences, and therefore having the same absolute time after pupal formation does not guarantee that pupae are at the same developmental stage. In many cases, stages defined by morphological features are preferable for comparing multiple individuals. Especially, a comparison between species requires precise staging and comparison among corresponding (homologous) stages.
Bainbridge and Bownes17 recognized 20 pupal stages (P1 to P15(ii)) based on morphological features of Drosophila melanogaster pupae. This staging is the most widely used system of morphological developmental staging18. In a previous study, we performed pupal staging of Drosophila guttifera to establish a basis for wing pigmentation studies19. D. guttifera has a black polka-dot pattern on its wings and is one of the model species for wing pigmentation formation20. Although we referred to the morphological criteria described in the Bainbridge and Bownes&#39; research17, we directly measured stage durations by serial observations19, instead of using Bainbridge and Bownes&#39; estimation of stage durations from observed frequency. Here we describe the method of pupal staging and measurement of durations of pupal stages of Drosophila used in Fukutomi et al19.
To study the developmental mechanism of wing pigmentation, we need to know when in pupal or adult stages the pigmentation occurs. Fukutomi et al.19 quantified optical densities (ODs) of pigmentation during pupal and adult stages by image analysis of wing images. The pigmentation of Drosophila wings is thought to be caused by accumulation of black melanin21. For quantification of ODs, gray-scale images and ImageJ software (https://imagej.nih.gov/ij/)22 were used. To recognize and quantify the spot-specific pigmentation (&#916;OD), we subtract the OD outside of a spot from the OD inside of a spot. To make this method reproducible and objective, the places of OD measurement should be determined using wing veins as landmarks. In this article, we describe in detail this method of quantification of wing pigmentation in Drosophila guttifera.

<table><tbody><tr><td>&lt;em&gt;Drosophila guttifera&lt;/em&gt;</td><td>The Drosophila Species Stock Center at the U.C. San Diego</td><td>15130-1971.10</td><td>&lt;em&gt;Drosophila guttifera&lt;/em&gt;, a fruit fly species used in this article</td></tr><tr><td>Plastic vial</td><td>Hightech</td><td>MKC-30</td><td>Plastic vial, for fly stock maintenance</td></tr><tr><td>Buzz plugs vial and bottle closures for glass vials</td><td>Fisher Scientific</td><td>AS-271</td><td>Cellulose plug, for fly stock maintenance</td></tr><tr><td>White soft sugar</td><td>Mitsui Sugar</td><td>J-500g</td><td>White soft sugar, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Corn flour</td><td>Nippon Flour Mills</td><td>F</td><td>Corn flour, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Corn grits - C</td><td>Nippon Flour Mills</td><td>GC</td><td>Corn grits - C, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Agar powder</td><td>Matsuki Kanten Sangyo</td><td>No.602</td><td>Agar powder, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Dry beer yeast</td><td>Asahi Food &amp;amp; Healthcare</td><td>Y2A</td><td>Dry beer yeast, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Butyl p-hydroxybenzoate</td><td>Nacalai Tesque</td><td>06327-02</td><td>Butyl p-hydroxybenzoate, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Ethanol</td><td>Wako</td><td>057-00456</td><td>Ethanol, for standard cornmeal/sugar/yeast/agar food</td></tr><tr><td>Flat bottom microtube</td><td>Ina Optica</td><td>CF-0150</td><td>1.5 mL microtube, for collecting pupae</td></tr><tr><td>CAPSULEFUGE</td><td>Tomy</td><td>PMC-060</td><td>Mini microcentrifuge, for collecting pupae</td></tr><tr><td>Sterilized Schale NB</td><td>Sansei Medical</td><td>01-013</td><td>Plastic Petri dish (diameter 90 mm x height 15 mm)</td></tr><tr><td>Serum tube rack</td><td>Iwaki</td><td>9796-050</td><td>Used as a moist chamber, for observation of pupa</td></tr><tr><td>Corning Falcon Easy-Grip tissue culture dish</td><td>Corning</td><td>353001</td><td>Plastic Petri dish (diameter 35 mm x height 10 mm)</td></tr><tr><td>Falcon standard tissue culture dish</td><td>Corning</td><td>353002</td><td>Plastic Petri dish (diameter 60 mm x height 15 mm)</td></tr><tr><td>Push-pin</td><td>Kokuyo</td><td>51233709</td><td>Push-pin, for making pinholes on the microtube lid</td></tr><tr><td>Stereomicroscope</td><td>Olympus</td><td>SZX16</td><td>Stereomicroscope, for morphological observation</td></tr><tr><td>Digital camera</td><td>Olympus</td><td>DSE-330-A</td><td>Digital camera, for imaging</td></tr><tr><td>NICETACK double sided tape</td><td>Nichiban</td><td>NW-15SF</td><td>Double sided tape, for removing puparium</td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tools</td><td>11252-20</td><td>Forceps, for removing puparium</td></tr><tr><td>Van Gogh VISUAL Paint brush</td><td>Talens Japan</td><td>GWVR-#5/0</td><td>Paint brush, for removing puparium</td></tr><tr><td>Greiner CELLSTAR 12 well cell culture plate</td><td>Merck</td><td>665-180</td><td>12-well cell culture plate, for measuring durations of pupal periods</td></tr><tr><td>NaCl</td><td>Wako</td><td>191-01665</td><td>NaCl, for PBS</td></tr><tr><td>KCl</td><td>Nacalai Tesque</td><td>285-14</td><td>KCl, for PBS</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;12H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Wako</td><td>196-02835</td><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;12H&lt;sub&gt;2&lt;/sub&gt;O, for PBS</td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Nacalai Tesque</td><td>28721-55</td><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, for PBS</td></tr><tr><td>Stepped Neutral Density (ND) Filter 0.04 - 3.0</td><td>Edmund Optics</td><td>64-384</td><td>Stepped density filter, for calibration of pigmentation measurement</td></tr><tr><td>ImageJ software</td><td>NIH</td><td>1.8.0-101</td><td>ImageJ software, for measurement of intensity of black spots on a wing (https://imagej.nih.gov)</td></tr><tr><td>FINE FROST glass slide</td><td>Matsunami Glass Ind</td><td>FF-001</td><td>Glass slide, for measurement of intensity of black spots on a wing</td></tr><tr><td>Square microscope cover glass 18 x 18</td><td>Matsunami Glass Ind</td><td>C018181</td><td>Cover slip, for measurement of intensity of black spots on a wing</td></tr></tbody></table>

methods for staging pupal periods and measurement of wing pigmentation of drosophila guttifera

Diversified species of Drosophila (fruit fly) provide opportunities to study mechanisms of development and genetic changes responsible for evolutionary changes. In particular, the adult stage is a rich source of morphological traits for interspecific comparison, including wing pigmentation comparison. To study developmental differences among species, detailed observation and appropriate staging are required for precise comparison. Here we describe protocols for staging of pupal periods and quantification of wing pigmentation in a polka-dotted fruit fly, Drosophila guttifera. First, we describe the method for detailed morphological observation and definition of pupal stages based on morphologies. This method includes a technique for removing the puparium, which is the outer chitinous case of the pupa, to enable detailed observation of pupal morphologies. Second, we describe the method for measuring the duration of defined pupal stages. Finally, we describe the method for quantification of wing pigmentation based on image analysis using digital images and ImageJ software. With these methods, we can establish a solid basis for comparing developmental processes of adult traits during pupal stages.

The pupal period of D. guttifera is divided into 17 stages (P1 - P15(ii); images of three representative stages (P1, P5 - 6, P10) are shown Figure 3, and all 17 stages are illustrated in Figure 4). Although Bainbridge and Bownes17 recognized 20 stages in D. melanogaster, some of these stages could not be applied to D. guttifera. The order of two developmental events, the appearan...

Watch this Scientific Journal Video about Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera at JoVE.com

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Protocols for staging pupal periods and measurement of wing pigmentation of Drosophila guttifera are described. Staging and quantification of pigmentation provide a solid basis for studying developmental mechanisms of adult traits and enable interspecific comparison of trait development.

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Diversified species of Drosophila (fruit fly) provide opportunities to study mechanisms of development and genetic changes responsible for evolutionary ...

methods-for-staging-pupal-periods-measurement-wing-pigmentation

Research

JoVE Journal

Biology

11.5K Views.  Kyoto University. Protocols for staging pupal periods and measurement of wing pigmentation of Drosophila guttifera are described. Staging and quantification of pigmentation provide a solid basis for studying developmental mechanisms of adult traits and enable interspecific comparison of trait development.

Video: Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

Wing development in Drosophila melanogaster is an ideal model for studying morphogenesis at tissue level. These appendages develop from a group of cells named wing imaginal discs formed during embryonic development. In the larval stages the imaginal discs grow, increasing its number of cells and forming monolayered epithelial&#160;structures. Inside the pupal case, the imaginal discs bud out and fold into bilayers along a line that becomes the future margin of the wing. During this process, the longitudinal primodia veins originate vein cells on the prospective dorsal and ventral surfaces of the wing. During the pupal stage the stripes of vein cells of each surface communicate in order to generate tight tubes; at the same time, the cross-veins begin their formation.
With the help of appropriate molecular markers, it is possible to identify the major elements composing the wing during its development. For this reason, the ability to accurately detect transcripts or proteins in this structure is critical for studying their abundance and localization related to the development process of the wing.
The procedure described here focuses on manipulating pupal wings, providing detailed instructions on how to dissect the wing during the pupal stage. The dissection of pupal tissue is more difficult to perform than their counterparts in third instar larvae. This is why this approach was developed, to obtain rapid and efficient high quality samples. Details of how to immunostain and mount these wing samples, to allow the visualization of proteins or cell components, are provided in the protocol. With little expertise it is possible to collect 8-10 high quality pupal wings in a short amount of time.

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

The ability to accurately detect transcripts or proteins in Drosophila tissues is critical for studying their abundance and localization related to the development process. Here is the description of a straightforward procedure to dissect pupal wings. These wings can be used as samples in several techniques (immunohistochemistry, PCR assay, etc.).

Wing development in Drosophila melanogaster is an ideal model for studying morphogenesis at tissue level. These appendages develop from a group of cells ...

Developmental Biology

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other insects proprioception is provided by specialized sensory organs termed chordotonal organs (ChOs) 2. Like many other organs in Drosophila, ChOs develop twice during the life cycle of the fly. First, the larval ChOs develop during embryogenesis. Then, the adult ChOs start to develop in the larval imaginal discs and continue to differentiate during metamorphosis.
The development of larval ChOs during embryogenesis has been studied extensively 10,11,13,15,16. The centerpiece of each ChO is a sensory unit composed of a neuron and a scolopale cell. The sensory unit is stretched between two types of accessory cells that attach to the cuticle via specialized epidermal attachment cells 1,9,14. When a fly larva moves, the relative displacement of the epidermal attachment cells leads to stretching of the sensory unit and consequent opening of specific transient receptor potential vanilloid (TRPV) channels at the outer segment of the dendrite 8,12. The elicited signal is then transferred to the locomotor central pattern generator circuit in the central nervous system.
Multiple ChOs have been described in the adult fly 7. These are located near the joints of the adult fly appendages (legs, wings and halters) and in the thorax and abdomen. In addition, several hundreds of ChOs collectively form the Johnston's organ in the adult antenna that transduce acoustic to mechanical energy 3,5,17,4.
In contrast to the extensive knowledge about the development of ChOs in embryonic stages, very little is known about the morphology of these organs during larval stages. Moreover, with the exception of femoral ChOs 18 and Johnston's organ, our knowledge about the development and structure of ChOs in the adult fly is very fragmentary.
Here we describe a method for staining and visualizing ChOs in third instar larvae and pupae. This method can be applied together with genetic tools to better characterize the morphology and understand the development of the various ChOs in the fly.

Visualization of Proprioceptors in Drosophila Larvae and Pupae

A method to immunostain and visualize chordotonal organs in larvae and pupae of Drosophila melanogaster is described.

Proprioception is the ability to sense the motion, or position, of body parts by responding to stimuli arising within the body. In fruitflies and other ...

Neuroscience

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Experimental Manipulation of Body Size to Estimate Morphological Scaling Relationships in Drosophila

The scaling of body parts is a central feature of animal morphology1-7. Within species, morphological traits need to be correctly proportioned to the body for the organism to function; larger individuals typically have larger body parts and smaller individuals generally have smaller body parts, such that overall body shape is maintained across a range of adult body sizes. The requirement for correct proportions means that individuals within species usually exhibit low variation in relative trait size. In contrast, relative trait size can vary dramatically among species and is a primary mechanism by which morphological diversity is produced. Over a century of comparative work has established these intra- and interspecific patterns3,4.
Perhaps the most widely used approach to describe this variation is to calculate the scaling relationship between the size of two morphological traits using the allometric equation y=bx&alpha;, where x and y are the size of the two traits, such as organ and body size8,9. This equation describes the within-group (e.g., species, population) scaling relationship between two traits as both vary in size. Log-transformation of this equation produces a simple linear equation, log(y) = log(b) + &alpha;log(x) and log-log plots of the size of different traits among individuals of the same species typically reveal linear scaling with an intercept of log(b) and a slope of &alpha;, called the 'allometric coefficient'9,10. Morphological variation among groups is described by differences in scaling relationship intercepts or slopes for a given trait pair. Consequently, variation in the parameters of the allometric equation (b and &alpha;) elegantly describes the shape variation captured in the relationship between organ and body size within and among biological groups (see 11,12).
Not all traits scale linearly with each other or with body size (e.g., 13,14) Hence, morphological scaling relationships are most informative when the data are taken from the full range of trait sizes. Here we describe how simple experimental manipulation of diet can be used to produce the full range of body size in insects. This permits an estimation of the full scaling relationship for any given pair of traits, allowing a complete description of how shape covaries with size and a robust comparison of scaling relationship parameters among biological groups. Although we focus on Drosophila, our methodology should be applicable to nearly any fully metamorphic insect.

Experimental Manipulation of Body Size to Estimate Morphological Scaling Relationships in Drosophila

Morphological scaling relationships capture and describe organismal shape. We present a method to measure morphological scaling relationships across the natural range of body sizes in fully metamorphic insects. Using a simple diet manipulation we increase the distribution of trait sizes, permitting the accurate description of how shape and size co-vary.

The scaling of body parts is a central feature of animal morphology1-7. Within species, morphological traits need to be correctly proportioned to the body ...

Extended Live Imaging of Drosophila Pupal Leg Morphogenesis Without Puparium

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Over the past decades, significant progress has been made in understanding the mechanisms of cell fate determination. However, the process by which fate-determined cells form three-dimensional organismal shapes remains unclear. Recent advances in confocal microscopy have facilitated efforts to observe cell dynamics during development through live imaging. The Drosophila melanogaster pupa is ideal for live imaging due to its immobility, transparent pupal cuticle, and the availability of fluorescent reporter lines. A primary challenge for imaging is the puparium, the cuticle surrounding the pupa, which obstructs optical imaging. While previous methods involved either partial or complete removal of the puparium, maintaining pupal viability for extended periods after this procedure has remained challenging. Here, a simple method is presented for days-long live imaging of the Drosophila leg during the pupal stage, involving complete puparium removal. The method includes removing the puparium from a pupa adhered to double-sided tape, followed by assembling a small chamber on a glass-bottom dish to enclose the pupa and a drop of water. This setup is straightforward, reliable, and supports extended pupal survival by preventing desiccation. Long-term live imaging of the Drosophila pupa has significantly contributed to capturing how the adult leg undergoes dramatic three-dimensional structural changes over 2-3 days. These changes include the transient formation of an intriguing structure (the Parthenon-like structure) by epithelial cells, rapid tissue narrowing, joint formation, and bristle elongation. This method is applicable to the observation of various tissues and can potentially be combined with other techniques, such as optical gene induction, to advance the understanding of cell dynamics during the final shape formation of tissues in the pupal stage.

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Here, a protocol is presented for long-term live imaging of the Drosophila leg during the pupal stage, involving complete removal of the puparium. This protocol can also be applied to other tissues.

Over the past decades, significant progress has been made in understanding the mechanisms of cell fate determination. However, the process by which ...

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

During the rapid inflammatory response to tissue damage, cells of the innate immune system are quickly recruited to the injury site. Once at the wound, innate immune cells perform a number of essential functions, such as fighting infection, clearing necrotic debris, and stimulating matrix deposition. In order to fully understand the diverse signaling events that regulate this immune response, it is crucial to observe the complex behaviors of (and interactions that occur between) multiple cell lineages in vivo, and in real-time, with the high spatio-temporal resolution. The optical translucency and the genetic tractability of Drosophila embryos have established Drosophila as an invaluable model to live-image and dissect fundamental aspects of inflammatory cell behavior, including mechanisms of developmental dispersal, clearance of apoptotic corpses&#160;and/or microbial pathogens, and recruitment to wounds. However, more recent work has now demonstrated that employing a much later stage in the Drosophila lifecycle &#8211; the Drosophila pupa &#8211; offers a number of distinct advantages, including improved RNAi efficiency, longer imaging periods, and significantly greater immune cell numbers. Here we describe a protocol for imaging wound repair and the associated inflammatory response at the high spatio-temporal resolution in live Drosophila pupae. To follow the dynamics of both re-epithelialization and inflammation, we use a number of specific in vivo fluorescent markers for both the epithelium and innate immune cells. We also demonstrate the effectiveness of photo-convertible fluorophores, such as Kaede, for following the specific immune cell subsets, to track their behavior as they migrate to, and resolve from, the injury site.

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

Here we present a protocol for live-imaging wound repair and the associated inflammatory response at high spatio-temporal resolution in vivo. This method utilizes the pupal stage of Drosophila development to enable long-term imaging and tracking of specific cell populations over time and is compatible with efficient RNAi-mediated gene inactivation.

During the rapid inflammatory response to tissue damage, cells of the innate immune system are quickly recruited to the injury site. Once at the wound, ...

Immunology and Infection

Quantitative Comparison of cis-Regulatory Element (CRE) Activities in Transgenic Drosophila melanogaster

Gene expression patterns are specified by cis-regulatory element (CRE) sequences, which are also called enhancers or cis-regulatory modules. A typical CRE possesses an arrangement of binding sites for several transcription factor proteins that confer a regulatory logic specifying when, where, and at what level the regulated gene(s) is expressed. The full set of CREs within an animal genome encodes the organism&prime;s program for development1, and empirical as well as theoretical studies indicate that mutations in CREs played a prominent role in morphological evolution2-4. Moreover, human genome wide association studies indicate that genetic variation in CREs contribute substantially to phenotypic variation5,6. Thus, understanding regulatory logic and how mutations affect such logic is a central goal of genetics. 
Reporter transgenes provide a powerful method to study the in vivo function of CREs. Here a known or suspected CRE sequence is coupled to heterologous promoter and coding sequences for a reporter gene encoding an easily observable protein product. When a reporter transgene is inserted into a host organism, the CRE&prime;s activity becomes visible in the form of the encoded reporter protein. P-element mediated transgenesis in the fruit fly species Drosophila (D.) melanogaster7 has been used for decades to introduce reporter transgenes into this model organism, though the genomic placement of transgenes is random. Hence, reporter gene activity is strongly influenced by the local chromatin and gene environment, limiting CRE comparisons to being qualitative. In recent years, the phiC31 based integration system was adapted for use in D. melanogaster to insert transgenes into specific genome landing sites8-10. This capability has made the quantitative measurement of gene and, relevant here, CRE activity11-13 feasible. The production of transgenic fruit flies can be outsourced, including phiC31-based integration, eliminating the need to purchase expensive equipment and/or have proficiency at specialized transgene injection protocols. 
Here, we present a general protocol to quantitatively evaluate a CRE&prime;s activity, and show how this approach can be used to measure the effects of an introduced mutation on a CRE&prime;s activity and to compare the activities of orthologous CREs. Although the examples given are for a CRE active during fruit fly metamorphosis, the approach can be applied to other developmental stages, fruit fly species, or model organisms. Ultimately, a more widespread use of this approach to study CREs should advance an understanding of regulatory logic and how logic can vary and evolve.

Quantitative Comparison of cis-Regulatory Element CRE Activities in Transgenic Drosophila melanogaster

Phenotypic variation for traits can result from mutations in cis-regulatory element (CRE) sequences that control gene expression patterns. Methods derived for use in Drosophila melanogaster can quantitatively compare the levels of spatial and temporal patterns of gene expression mediated by modified or naturally occurring CRE variants.

Gene expression patterns are specified by cis-regulatory element (CRE) sequences, which are also called enhancers or cis-regulatory modules. A typical CRE ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Insect Puparium Removal: A Method for Pupal Investigation

Source: Fukutomi, Y., et al. <a href="https://www.jove.com/video/56935/methods-for-staging-pupal-periods-measurement-wing-pigmentation">Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of&#160;Drosophila guttifera.</a>&#160;J. Vis. Exp. (2018).
This article describes a technique to remove an insect&#8217;s puparium while keeping the inside pupa intact and therefore fit for further investigation. The example protocol shows the procedure with Drosophila guttifera; however, this technique is applicable to other Drosophila species.

Source: Fukutomi, Y., et al. Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of&#160;Drosophila guttifera.&#160;J. Vis. Exp. ...

Methods for Staging Pupal Periods and Measurement of Wing Pigmentation of Drosophila guttifera

Summary

Explore More Videos

Chapters in this video

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

Visualization of Proprioceptors in Drosophila Larvae and Pupae

Dissection and Staining of Drosophila Pupal Ovaries

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Experimental Manipulation of Body Size to Estimate Morphological Scaling Relationships in Drosophila

Long-Term Live Imaging of Drosophila Pupal Leg Development After Puparium Removal

Long-term In Vivo Tracking of Inflammatory Cell Dynamics Within Drosophila Pupae

Quantitative Comparison of cis-Regulatory Element (CRE) Activities in Transgenic Drosophila melanogaster

Imaging Through the Pupal Case of Drosophila melanogaster

Insect Puparium Removal: A Method for Pupal Investigation