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

Summary

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.

Abstract

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.

Introduction

Some of the morphological traits of Drosophila are diversified among species^1,2,3,4,5. 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 pigmentation^{6,7,8,9,10,11,12,13,14,15}. 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, "hours APF" (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' research¹⁷, we directly measured stage durations by serial observations¹⁹, instead of using Bainbridge and Bownes' 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 (ΔOD), 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.

Protocol

1. Fly stock

1. Use Drosophila guttifera for all of the following protocols.
2. 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.

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 °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.

1. 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.
2. 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.
3. Centrifuge the microtubes with food for 7 s in a mini microcentrifuge (860 x g).
4. Invert and tap vials to remove all adults from the vial.
5. Pour 5 - 10 mL of ddH₂O (or reverse osmosis water) into the vial.
6. 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).
7. Gently move late 3rd instar larvae with forceps into the microtubes with food (10 larvae / microtube). Incubate them overnight at 25 °C.
8. 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).
9. 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.
10. 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).
11. Observe pupae under a stereo microscope. Take photographs using a digital camera attached to the stereo microscope.

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.

1. Affix a piece of double-sided tape on a piece of paper towel.
2. Place a pupa on the double-sided tape ventral side up (Figure 1A).
3. 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.
4. 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).
5. After breaking the puparium as much as possible, take out the pupa using a fine paintbrush (#5/0) (Figure 1C).
6. 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.

4. Measuring durations of pupal stages

1. 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 °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.
2. 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 references^17,19) in a tally sheet.
3. Continue recording over four straight days (96 h) by a rotating shift of three (or more) persons.
4. 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).

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 (ΔOD). Here, we describe the method of dissection, measurement and calculation of ΔOD. 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.

1. Image preparation
   1. 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).
   2. 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).
   3. 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.
   4. Place 10 µL of PBS on a glass slide, place the wing there, and cover with a cover slip (18 mm x 18 mm).
   5. 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.
   6. 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.
2. Calibration
   1. Take images of 9 parts of a stepped density filter using the same camera settings used to obtain the wing image.
   2. Initiate ImageJ software (https://imagej.nih.gov/ij/)²².
   3. Click File | Open | and select one of the images of the stepped density filter.
   4. Click Image | Type | 8-bit | to convert the image to an 8-bit image.
   5. 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.
   6. Click Analyze | Measure|. The "mean grey value" of selected areas are measured.
   7. Repeat 5.2.3. to 5.2.6. for the 8 remaining images.
   8. 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).
   9. Check Global calibration column and click OK.
      NOTE: By performing this procedure, "mean grey value" is converted to "optical density (OD)" using the Rodbard function. After this step, optical density can be calculated for a particular selected area in ImageJ software.
3. 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.
   1. Definition of Point A, campaniform sensillum spot.
      1. 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.
      2. 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.
      3. Click Edit | Selection | Add to Manager |. Check Show All column.
      4. 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 "m" 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.
      5. Click Edit | Selection | Add to Manager |.
      6. 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.
      7. 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).
      8. Record the x coordinate and the y coordinate of Point A, indicated below Area Selection Tools when placing the cursor on Point A.
   2. Definition of Point B, longitudinal vein tip spot
      1. 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.
      2. Repeat the same procedure described in Step 5.3.1. (Definition of Point A, campaniform sensillum spot) to find Point A in the image.
      3. Click Edit | Selection | Add to Manager |.
      4. 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.
      5. Click Edit | Selection | Add to Manager |.
      6. 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 "m" key to measure the angle between the two lines.
      7. Click Edit | Selection | Add to Manager |.
      8. 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.
      9. Click Edit | Selection | Add to Manager |.
      10. 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).
      11. Record the x coordinate and the y coordinate of Point B, indicated below Area Selection Tools when placing the cursor on Point B.
   3. Definition of Point C, posterior crossvein spot
      1. 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.
      2. 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).
      3. Record the x coordinate and the y coordinate of Point C, indicated below Area Selection Tools when placing the cursor on Point C.
   4. Definition of Point D, control area
      1. 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.
      2. 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).
      3. Record the x coordinate and the y coordinate of Point D, indicated below Area Selection Tools when placing the cursor on Point D.
4. Measurements
   1. 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.
   2. Click Rectangle in Area Selection Tools and draw a rectangle of approximately 1/9 size of the image window.
   3. 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.
   4. Click Analyze | Measure |. If the calibration described in 5.2 has already been finished, ODs are indicated in Mean column.
   5. Calculate ΔODs 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.

Results

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...

Discussion

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...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Drosophila guttifera	The Drosophila Species Stock Center at the U.C. San Diego	15130-1971.10	Drosophila guttifera, a fruit fly species used in this article
Plastic vial	Hightech	MKC-30	Plastic vial, for fly stock maintenance
Buzz plugs vial and bottle closures for glass vials	Fisher Scientific	AS-271	Cellulose plug, for fly stock maintenance
White soft sugar	Mitsui Sugar	J-500g	White soft sugar, for standard cornmeal/sugar/yeast/agar food
Corn flour	Nippon Flour Mills	F	Corn flour, for standard cornmeal/sugar/yeast/agar food
Corn grits - C	Nippon Flour Mills	GC	Corn grits - C, for standard cornmeal/sugar/yeast/agar food
Agar powder	Matsuki Kanten Sangyo	No.602	Agar powder, for standard cornmeal/sugar/yeast/agar food
Dry beer yeast	Asahi Food & Healthcare	Y2A	Dry beer yeast, for standard cornmeal/sugar/yeast/agar food
Butyl p-hydroxybenzoate	Nacalai Tesque	06327-02	Butyl p-hydroxybenzoate, for standard cornmeal/sugar/yeast/agar food
Ethanol	Wako	057-00456	Ethanol, for standard cornmeal/sugar/yeast/agar food
Flat bottom microtube	Ina Optica	CF-0150	1.5 mL microtube, for collecting pupae
CAPSULEFUGE	Tomy	PMC-060	Mini microcentrifuge, for collecting pupae
Sterilized Schale NB	Sansei Medical	01-013	Plastic Petri dish (diameter 90 mm x height 15 mm)
Serum tube rack	Iwaki	9796-050	Used as a moist chamber, for observation of pupa
Corning Falcon Easy-Grip tissue culture dish	Corning	353001	Plastic Petri dish (diameter 35 mm x height 10 mm)
Falcon standard tissue culture dish	Corning	353002	Plastic Petri dish (diameter 60 mm x height 15 mm)
Push-pin	Kokuyo	51233709	Push-pin, for making pinholes on the microtube lid
Stereomicroscope	Olympus	SZX16	Stereomicroscope, for morphological observation
Digital camera	Olympus	DSE-330-A	Digital camera, for imaging
NICETACK double sided tape	Nichiban	NW-15SF	Double sided tape, for removing puparium
Dumont #5 forceps	Fine Science Tools	11252-20	Forceps, for removing puparium
Van Gogh VISUAL Paint brush	Talens Japan	GWVR-#5/0	Paint brush, for removing puparium
Greiner CELLSTAR 12 well cell culture plate	Merck	665-180	12-well cell culture plate, for measuring durations of pupal periods
NaCl	Wako	191-01665	NaCl, for PBS
KCl	Nacalai Tesque	285-14	KCl, for PBS
Na2HPO4·12H2O	Wako	196-02835	Na2HPO4·12H2O, for PBS
KH2PO4	Nacalai Tesque	28721-55	KH2PO4, for PBS
Stepped Neutral Density (ND) Filter 0.04 - 3.0	Edmund Optics	64-384	Stepped density filter, for calibration of pigmentation measurement
ImageJ software	NIH	1.8.0-101	ImageJ software, for measurement of intensity of black spots on a wing (https://imagej.nih.gov)
FINE FROST glass slide	Matsunami Glass Ind	FF-001	Glass slide, for measurement of intensity of black spots on a wing
Square microscope cover glass 18 x 18	Matsunami Glass Ind	C018181	Cover slip, for measurement of intensity of black spots on a wing