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>

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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>Drosophila guttifera</td>
			<td>The Drosophila Species Stock Center at the U.C. San Diego</td>
			<td>15130-1971.10</td>
			<td>Drosophila guttifera, 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 &#38; 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>Na2HPO4&#183;12H2O</td>
			<td>Wako</td>
			<td>196-02835</td>
			<td>Na2HPO4&#183;12H2O, for PBS</td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Nacalai Tesque</td>
			<td>28721-55</td>
			<td>KH2PO4, 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 ...

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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

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP&#8217;s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP&#8217;s and minature excitatory junction potentials (mEJP&#8217;s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2 (HL3) that contains 5 mM Mg2+ and 1.5 mM Ca2+. Changes in the amplitude of evoked EJP&#8217;s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.

Here we describe electrophysiological methods for measuring synaptic transmission at the neuromuscular junction of Drosophila larva. Evoked release is initiated artificially by stimulating the motor neuron axons, and transmission through the NMJ can be measured by the postsynaptic response evoked in the muscle.

In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. ...

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations. In these preparations, the central nervous system runs down the middle, and is flanked by the body walls. Many different phenotypes have been examined using such preparations. In most cases, the fillets were generated by dissection of antibody-stained fixed whole-mount embryos. These "fixed dissections" have some disadvantages, however. They are time-consuming to execute, and it is difficult to sort mutant (GFP-negative) embryos from stocks in which mutations are maintained over GFP balancer chromosomes. Since 2002, our group has been conducting deficiency and ectopic expression screens to identify ligands for orphan receptors. In order to do this, we developed streamlined protocols for live embryo dissection and antibody staining of collections containing hundreds of balanced lines. We have concluded that it is considerably more efficient to examine phenotypes in large collections of stocks by live dissection than by fixed dissection. Using the protocol described here, a single trained individual can screen up to 10 lines per day for phenotypes, examining 4-7 mutant embryos from each line under a compound microscope. This allows the identification of mutations conferring subtle, low-penetrance phenotypes, since up to 70 hemisegments per line are scored at high magnification with a 40X water-immersion lens.

Live Dissection of Drosophila Embryos: Streamlined Methods for Screening Mutant Collections by Antibody Staining

We describe a streamlined protocol for generating "fillet" preparations of Drosophila embryos of specific genotypes. This protocol allows efficient execution of a variety of genetic screens. It also allows excellent visualization of structures in the late embryo.

Drosophila embryos between stages 14 and 17 of embryonic development can be readily dissected to generate "fillet" preparations.  In these preparations, ...

Laser Microdissection Applied to Gene Expression Profiling of Subset of Cells from the Drosophila Wing Disc

Heterogeneous nature of tissues has proven to be a limiting factor in the amount of information that can be generated from biological samples, compromising downstream analyses. Considering the complex and dynamic cellular associations existing within many tissues, in order to recapitulate the in vivo interactions thorough molecular analysis one must be able to analyze specific cell populations within their native context. Laser-mediated microdissection can achieve this goal, allowing unambiguous identification and successful harvest of cells of interest under direct microscopic visualization while maintaining molecular integrity. We have applied this technology to analyse gene expression within defined areas of the developing Drosophila wing disc, which represents an advantageous model system to study growth control, cell differentiation and organogenesis. Larval imaginal discs are precociously subdivided into anterior and posterior, dorsal and ventral compartments by lineage restriction boundaries. Making use of the inducible GAL4-UAS binary expression system, each of these compartments can be specifically labelled in transgenic flies expressing an UAS-GFP transgene under the control of the appropriate GAL4-driver construct. In the transgenic discs, gene expression profiling of discrete subsets of cells can precisely be determined after laser-mediated microdissection, using the fluorescent GFP signal to guide laser cut. 
Among the variety of downstream applications, we focused on RNA transcript profiling after localised RNA interference (RNAi). With the advent of RNAi technology, GFP labelling can be coupled with localised knockdown of a given gene, allowing to determinate the transcriptional response of a discrete cell population to the specific gene silencing. To validate this approach, we dissected equivalent areas of the disc from the posterior (labelled by GFP expression), and the anterior (unlabelled) compartment upon regional silencing in the P compartment of an otherwise ubiquitously expressed gene. RNA was extracted from microdissected silenced and unsilenced areas and comparative gene expression profiling determined by quantitative real-time RT-PCR. We show that this method can effectively be applied for accurate transcriptomics of subsets of cells within the Drosophila imaginal discs. Indeed, while massive disc preparation as source of RNA generally assumes cell homogeneity, it is well known that transcriptional expression can vary greatly within these structures in consequence of positional information. Using localized fluorescent GFP signal to guide laser cut, more accurate transcriptional analyses can be performed and profitably applied to disparate applications, including transcript profiling of distinct cell lineages within their native context.

Laser Microdissection Applied to Gene Expression Profiling of Subset of Cells from the Drosophila Wing Disc

Laser microdissection was applied to analyse gene expression profiling in specific compartments of Drosophila wing disc subjected to localised RNAi in vivo. RNA extracted from equivalent areas of silenced and unsilenced compartments was analysed by quantitative RT-PCR to determine comparative gene expression profiling within the context of native tissue microecology.

Heterogeneous nature of tissues has proven to be a limiting factor in the amount of information that can be generated from biological samples, ...

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Drosophila has proven to be a useful model system for analysis of behavior, including flight. The initial flight tester involved dropping flies into an oil-coated graduated cylinder; landing height provided a measure of flight performance by assessing how far flies will fall before producing enough thrust to make contact with the wall of the cylinder. Here we describe an updated version of the flight tester with four major improvements. First, we added a &#34;drop tube&#34; to ensure that all flies enter the flight cylinder at a similar velocity between trials, eliminating variability between users. Second, we replaced the oil coating with removable plastic sheets coated in Tangle-Trap, an adhesive designed to capture live insects. Third, we use a longer cylinder to enable more accurate discrimination of flight ability. Fourth we use a digital camera and imaging software to automate the scoring of flight performance. These improvements allow for the rapid, quantitative assessment of flight behavior, useful for large datasets and large-scale genetic screens.

An Improved Method for Accurate and Rapid Measurement of Flight Performance in Drosophila

Here we describe a method for rapid and accurate measurement of flight performance in Drosophila, enabling high-throughput screening.

Drosophila has proven to be a useful model system for analysis of behavior, including flight. The initial flight tester involved dropping flies into an ...

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 ...

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial ...

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes differentiation from a stem cell population, mitotic amplification, and meiosis. In addition, post-meiotic germ cells undergo a dramatic morphological reshaping process as well as a global epigenetic reconfiguration of the germ line chromatin&#8212;the histone-to-protamine switch.
Studying the role of a protein in post-meiotic spermatogenesis using mutagenesis or other genetic tools is often impeded by essential embryonic, pre-meiotic, or meiotic functions of the protein under investigation. The post-meiotic phenotype of a mutant of such a protein could be obscured through an earlier developmental block, or the interpretation of the phenotype could be complicated. The model organism Drosophila melanogaster offers a bypass to this problem: intact testes and even cysts of germ cells dissected from early pupae are able to develop ex vivo in culture medium. Making use of such cultures allows microscopic imaging of living germ cells in testes and of germ-line cysts. Importantly, the cultivated testes and germ cells also become accessible to pharmacological inhibitors, thereby permitting manipulation of enzymatic functions during spermatogenesis, including post-meiotic stages.
The protocol presented describes how to dissect and cultivate pupal testes and germ-line cysts. Information on the development of pupal testes and culture conditions are provided alongside microscope imaging data of live testes and germ-line cysts in culture. We also describe a pharmacological assay to study post-meiotic spermatogenesis, exemplified by an assay targeting the histone-to-protamine switch using the histone acetyltransferase inhibitor anacardic acid. In principle, this cultivation method could be adapted to address many other research questions in pre- and post-meiotic spermatogenesis.

Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment

This protocol describes the dissection and cultivation of intact testes and germ-line cysts from Drosophila melanogaster pupae. This method allows microscopic observation of spermatogenesis ex vivo. Furthermore, we describe a pharmacological assay of the effect of inhibitors on specific stages of germ-cell development in pupal testes.

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes ...