Colorimetric synchronization of Drosophila larvae

Abstract

The rapid succession of events during development poses an inherent challenge to achieve precise synchronization required for rigorous, quantitative phenotypic and genotypic analyses in multicellular model organisms. Drosophila melanogaster is an indispensable model for studying the development and function of higher order organisms due to extensive genome homology, tractability, and its relatively short lifespan. Presently, nine Nobel prizes serve as a testament to the utility of this elegant model system. Ongoing advancements in genetic and molecular tools allow for the underlying mechanisms of human disease to be investigated in Drosophila. However, the absence of a method to precisely age-match tissues during larval development prevents further capitalization of this powerful model organism. Drosophila spend nearly half of their life cycle progressing through three morphologically distinct larval instar stages during which the imaginal discs, precursors of mature adult external structures (e.g., eyes, legs, wings), grow and develop distinct cell fates. Other tissues, such as the central nervous system, undergo massive morphological changes during larval development. While these three larval stages and subsequent pupal stages have historically been identified based on hours after egg-laying under standard laboratory conditions, a reproducible, efficient, and inexpensive method to accurately age-match larvae within the third instar is required. The third instar stage is of particular interest, as this developmental stage spans a 48-hour window during which larval tissues switch from proliferative to differentiation programs. Moreover, some genetic manipulations can lead to developmental delays, further compounding the need for precise age-matching between control and experimental samples. This article provides a protocol optimized for synchronous staging of Drosophila third instar larvae by colorimetric characterization and is useful for age-matching a variety of tissues for numerous downstream applications. We also provide a brief discussion of the technical challenges associated with successful application of this protocol.

Basic Protocol 1: Synchronization of third instar Drosophila larvae

INTRODUCTION:

As Drosophila progress through their approximately 11-day lifecycle, they develop from embryos to larvae to sexually dimorphic adults (Figure 1A). This metamorphosis transforms physical characteristics, including size, biological complexity, and behavior, and is further hallmarked by distinct changes in gene expression and hormone signaling. Dramatic reorganizations of tissues and organ systems are also noted throughout these developmental periods (Tissot & Stocker, 2000, Coronado-Zamora, 2019).

Figure 1. Lifespan of Drosophila and development of adult structures from larval imaginal discs.

(A) Diagram showing the progression of the Drosophila life cycle and typical developmental timing (25 °C). (B) Cartoon shows several color-coded primordial structures within a third instar larva and corresponding structures within an adult. Images were generated using BioRender.com.

One challenge to analyzing key mechanisms of development or disease involves accurately comparing tissue from similarly staged samples, as in a genetic control to a mutant, or a treated versus mock-treated control. Such studies require samples to be both chronologically and developmentally age-matched. Developmental synchronization ensures that studies are done on samples within the range of developmental events of interest, thereby enhancing precision and reproducibility.

The Drosophila third larval instar poses a unique challenge for synchronization, as this developmental period spans a 48-hour period marked by changes in cell fate, the onset of local patterning, and large-scale changes in tissue architecture. Many biological questions can be answered within third instar larvae, including how cell types are differentiated and serve as direct precursors to adult structures (Figure 1B). Thus, a technique is required to age-match third instar larvae. Historical methods to select age-matched larvae are labor-intensive, error-prone, and imprecise (Ashburner, 1967). To more precisely stage and select third instar larvae, we combined dual synchronization with colorimetric tracing. Age-matched samples can be further manipulated, dissected, fixed, and/or prepared for biochemical, genomic, or imaging analyses. This protocol is useful for the study of various mechanisms underlying regeneration, neurodevelopment, metabolism, germline formation, behavior, and numerous other biological processes.

The protocol relies upon the semi-transparent nature of Drosophila larval development to stage samples based on gut clearance of colored food. The introduction of a chemical dye to food enables investigators to track defined checkpoints through mid-late larval stages. This protocol is suited for experiments in which the purpose is to study events requiring sample-to-sample comparisons and to ensure samples within a given group are of comparable age. This protocol details successive synchronization steps commencing during embryogenesis and continuing through larval molting to yield samples that are paired in both chronological and developmental age.

CAUTION: Molten agar poses a burn hazard. Use care and appropriate personal protective equipment to avoid contact with skin.

BASIC PROTOCOL 1

Synchronization of third instar Drosophila larvae

Materials:

Grape agar plates (see recipe)

Yeast paste (see recipe)

Drosophila melanogaster (Bloomington Drosophila Resource Center)

Drosophila media in vials (Lab-Express, Inc., cat. no. 7001-WV)

Blue yeast paste (see recipe)

Embryo collection cage (Genesee Scientific, cat. no 59-101, or equivalent)

60 mm - petri dishes (Fisher Scientific, cat. no. FB0875713A)

Light and temperature-controlled incubator (Shel Labs or equivalent)

Stereo light microscope (Zeiss Stemi 508 or equivalent)

Anesthesia apparatus (e.g., carbon dioxide flypad)

Paint brush (Dick Blick, cat. no. 06376-7050; size 5/0, 1.5 mm nylon or equivalent)

Precision probe (Fisher Scientific, cat. no. 12-000-153)

½” - Laboratory tape (VWR, cat. no. 89097-916)

Spoonula lab spoon (Fisher Scientific, cat. no. S35888)

Protocol steps with step annotations:

Seed embryo collection cage

• 1Seed cage(s) with approximately 50–100 female and 20–30 male Drosophila.

  Genotypes of Drosophila used will vary depending on experimental design. Usually, freshly eclosed adults (0–3 days) are used to maximize fecundity, which decreases with age. Drosophila are sorted on the anesthesia apparatus using a paint brush. We use a custom-built 60 mm diameter acrylic cage sealed on one end with gas-permeable 30 μm Nitex mesh. Commercial embryo collection cages in 35-mm or 100-mm sizes are available from Genesee Scientific. Grape juice agar plates should be poured in an appropriately sized petri dish to ensure a snug fit with the embryo collection cage. We use 60 mm dishes to accommodate our 60 mm cages.

• 2Affix a grape juice agar plate supplemented with yeast paste to the open end of the cage using laboratory tape.

  Adjusting the water added to the grape agar mix will alter the consistency of the agar plate (see recipe). Premade agar plates may be stored at 4 °C for up to one month but should be pre-warmed to room temperature before use. Ensure proper fitting of the grape agar plate to avoid loss or contamination of Drosophila within the cage. For some incubators, a humidity source may be needed to prevent excessive drying of the agar plates. Avoid pushing the cage too hard onto the agar.

• 3Store seeded cage upright in light and temperature-controlled incubator at 25°C.

  An optional step is to cover the embryo collection cage with cheesecloth to further limit stray Drosophila entering the cage.

• 4Replace grape plate after 24 hours.

  Cages must be provided with fresh, yeasted, pre-warmed grape plates daily. Changing grape plates on the cage involves inverting the cage, carefully peeling back the tape while gently holding down the plate to prevent flies from escaping, tapping the cage down on a soft pad (e.g., a mouse pad), deftly removing the old plate and replacing it with a fresh one, retaping the plate, and returning the cage upright. Avoid use of anesthesia, which can alter downstream responses, including embryo yield. Animals will require between 24–48 hours to acclimate to the cage setting, during which egg laying may be sub-optimal.

Embryo collection

• 5Replace grape juice agar plate with a fresh, yeasted grape plate.
• 6Incubate cage at 25°C for 30–60 minutes.

  This step pre-clears embryos stored within mated females and is a key step for initially synchronizing the embryo collection.

• 7Replace grape juice agar plate with a fresh, yeasted grape juice agar plate.
• 8Incubate cage at 25°C for 4 hours.

  This step will yield a 0–4 hour synchronized embryo collection.

• 9Recover the seeded grape juice agar plate.
• 10Provide embryo collection cage with a fresh, yeasted grape plate if needed.

  If no further embryo collections are needed, the embryo collection cage may be incubated at −20°C ≥4-hours to kill remaining adult Drosophila. Follow institutional environmental health and safety office guidelines to dispose of fly remains responsibly. Cages may be washed and reused.

• 11Cover the seeded grape juice plate with a lid.
• 12Ensure the seeded plate is appropriately labeled.
• 13Incubate the seeded plate in the 25°C incubator for 24 hours.

  This incubation follows Drosophila embryogenesis to completion. Viable embryos will hatch into first instar larvae (Figure 1A). Developmental progression is temperature-sensitive and will vary based on genotype and/or sample conditions.

Larval transfer

• 14Score Drosophila culturing medium within a vial using the precision probe.
• 15Transfer a spoonful of blue yeast paste (see recipe) into the vial using a clean spoonula.

  Please note: Bromophenol blue is a pH indicator and therefore, should not be used in studies where the pH in the gut may change. Alternatively, a food-grade dye like brilliant blue FCF may be used.

• 16Remove seeded grape agar plate from the 25°C incubator, and place under the stereo light microscope.
• 17Use the precision probe to transfer first instar larvae into the culturing vial.

  Avoid over-manipulation of the larvae. The precision tool may be used to gently scoop up larvae. A one-quarter turn twisting motion may be useful to deposit larvae carefully on the Drosophila culturing medium surface. Avoid gouging larvae into the medium. Up to 100 larvae may be transferred to a single vial. Prepare additional vials, as needed.

• 18Incubate the seeded vial(s) in the 25°C incubator for 72–96 hours.

  This step synchronizes the start of larval development. During this period, Drosophila larvae will complete the first and second larval instars and enter the third larval instar stage (Figure 1A). Developmental progression is temperature-sensitive and may vary depending on genotype and/or sample conditions.

Colorimetric selection

• 19Confirm presence of third instar larvae by identification of foraging behavior.

  Wandering third instar larvae will freely migrate up the sides of the vial and explore their environment. Wandering behavior develops as the larvae stop feeding. During this period, the dyed food will gradually clear from the intestine.

• 20Select the desired third larval instar stage according to graded gut clearance of the blue yeast paste (Figure 2).

  Dark blue staining visible within the gut indicates early third instar larvae that may be expected to pupariate within 12–24 hours. Light or partial blue staining represents partial gut clearance; these larvae may pupariate within 5–12 hours. Complete gut clearance is marked by the absence of blue staining characteristic of late third instar larvae that may pupariate within 1–6 hours. Bromophenol blue dye has not been reported to elicit effects on animal development. Developmental timeframes vary depending on animal genotypes or experimental conditions.

Figure 2. The three stages of a third instar larva.

Cartoons show the passage of blue colored food through the larval intestinal tract. Beneath each schematic is a representative image of the corresponding stage from samples processed using this protocol. Bars: 1mm. Cartoons were created using BioRender.com.

REAGENTS AND SOLUTIONS:

Grape juice agar plates

• Grape agar plate premix (Genesee Scientific, cat. no. 47-102)

• 380 ml water

• Prepare by following the manufacturer’s instructions

• Use caution to avoid agar from boiling over

• Pour about 10 ml molten agar mix per 60 mm petri dish

• Store solidified plates inverted at 4°C for up to 1 month in a covered food storage container

Yeast paste

• 2 g active dry yeast (Genesee Scientific, cat. no. 62-103)

• 10 ml water

• Mix thoroughly with scoopula, adjusting volumes as needed to reach desired consistency of smooth peanut butter

• Store, covered at 4°C for up to 1 month

Blue yeast paste

• 1 g active dry yeast (Genesee Scientific, cat. no. 62-103)

• 5 ml water

• 0.05% w/v bromophenol blue (Fisher Scientific, cat. no. BP115-25)

• Mix thoroughly with scoopula, scaling volumes as needed

• Store, covered at 4°C for up to 1 month

COMMENTARY:

Background information

Drosophila melanogaster are a powerful and popular model system utilized to study fundamental biological processes and molecular mechanisms underlying human disease. Drosophila share significant functional and genetic homologies with humans and other higher-order organisms. Advantages of the Drosophila model include a shortened life cycle, ease of genetic and molecular manipulation, tractability for genetic and functional analyses, and cost-effectiveness (Pandey & Nichols, 2011).

As they develop between embryonic, larval, pupa, and adult stages, Drosophila transform numerous characteristics, including size, color, biological complexity, gene expression patterns, and behavior. The distinct morphological changes that mark Drosophila development are defined by the onset of developmental waves characterized by gene expression and signaling cascades (Coronado-Zamora, Salvador-Martínez, Castellano, Barbadilla, & Salazar-Ciudad, 2019). As holometabolous insects, Drosophila undergo a series of larval molts that define three morphologically distinct larval stages characterized by hormonal remodeling of many organ systems, including the nervous system (Tissot & Stocker, 2000). Many functional tissues in adult Drosophila are directly derived from progenitor tissues established during these larval stages, prompting researchers to prioritize sampling larvae to identify and characterize molecular pathways involved in tissue growth and patterning.

The three larval instars of Drosophila are distinguished based on size, behavior, duration, and developmental time, represented as hours after egg-laying (AEL) (Wegman, Ainsley, & Johnson, 2010). However, each larval stage persists for one or more days, necessitating more accurate methods to bin age-matched samples. For example, the third instar larval stage spans approximately 48-hours (Figure 1A). To examine responses within a single population or across conditions or genotypes, multiple samples must be cross-compared. Chronologic and developmental age-matching increases the precision of experimental and biological replicates to ensure observed variances are true effects, rather than artefacts introduced through invalid comparisons between different developmental stages.

Historical strategies to define larval stages are labor-intensive and require specialized training. The timing of salivary gland chromosome puffing activity, mean size, or both have been used to distinguish third instar larvae into defined puff stages (PS1–9). Polytene chromosomes found in the developing salivary gland nuclei permit visualization of active or repressed genes based on classically defined puff versus banding patterns (Beermann & Clever, 1964). The relative timeline of each larval stage may be assessed by analysis of chromosome organization or puffing stage at certain loci within salivary gland preparations (Ashburner, 1967). Within any larval stage, puffs can appear, regress, and even reappear under a strict chronological sequence, often spanning hours. Though this technique identifies larval stages, patterns between some stages, including late third instar larva versus early pupa, are difficult to distinguish. Additionally, variance in banding patterns can make comparing some genotypes or across species difficult (Ashburner, 1967; Rodman & Kopac, 1964).

Currently, the standard practice to stage third instar larvae relies on egg lay synchronization and/or behavioral observation (e.g., observing the wandering behavior of mid-late third instar larvae). An important limitation of the standard practice is the inability to efficiently forecast the number of similarly staged larvae that will develop within the next day. Therefore, a remaining gap is a method to more precisely age-match larval samples across genotypes, conditions, and experiments. An approach that could accurately age-match larvae at distinct stages within the third instar would increase reproducibility and rigor by increasing experimental sample sizes, leading to more biologically and developmentally relevant data, as well as increased efficiencies in experimental output.

Here we present a method that combines embryonic and early larval synchronization with the introduction of a traceable colorimetric marker to accurately stage third instar larvae by intestinal gut clearance. This approach has several advantages in that it does not compromise the behavior or viability of the samples, it is inexpensive, robust, and requires no specialized training or equipment beyond what is typically found in most Drosophila laboratories. This simple and low-cost technique is accessible to research laboratories in primarily undergraduate institutions or under-resourced countries.

With this technique, Drosophila development is synchronized first during embryogenesis, then again, at the first larval instar. First instar larvae are then cultured in tandem with a nutritive yeast paste containing the common pH indicator and tracing dye, bromophenol blue. The pigmentation of Drosophila larvae is relatively light and transparent, which renders the ingestion of bromophenol blue visible as a tracer of food intake and progression through the digestive tract (Andres & Thummel, 1994; Maroni, Laurie-Ahlberg, Adams, & Wilton, 1982)

A key principle of this protocol stems from the finding that food clearance tracks with third larval development (Andres & Thummel, 1994). Specifically, early third instar larvae selected using this technique will have a dark blue gut at the anterior and correspond to PS1. These animals will take 12–24 hours to pupariate. In contrast, mid- third instar larvae with partially cleared intestines (apparent by lighter-blue staining and reduced internalized dye) correspond to PS2–6 and will pupariate within 5–12 hours. Finally, late third instar larvae that correspond to PS7–9 will pupariate within 1–6 hours and are indicated by total gut clearance of the dye (Figure 2) (Andres & Thummel, 1994; Ashburner, 1967). Bromophenol blue is an acid-base indicator, which highlight a caveat if investigating a process that may affect the pH of the gut. This may confound developmental selection of samples, or otherwise make results difficult to interpret. An alternative dye may be used, such as brilliant blue FCF.

Larval synchronization is critical to ensure that samples are indeed age-matched within the ranges(s) of key developmental events and that observed differences are not due to indirect effects associated with inaccurate chronological age or developmental stage. This protocol presents an accessible technique to accurately synchronize Drosophila larvae beginning at the embryonic stage.

Critical Parameters:

Temperature, and more broadly, culturing conditions, largely influence this protocol. Special attention should be paid to house animals in a light and temperature-controlled environment, at indicated metrics. Drosophila specimens must be maintained at defined temperatures (25°C) to utilize the proposed timeline. We and others have adapted this protocol to accurately age-match samples following temperature-shift experiments, which will alter the rate of developmental progression. Any deviation in culture conditions will offset the staging of larval samples.

Removing and replacing grape juice agar plates must be done quickly. Novice users may benefit from practice to avoid loss of adult Drosophila from the cage. We avoid anaesthetizing animals during cage changes to prevent any downstream effects caused by anesthesia.

The transfer of first instar larvae from agar plates to Drosophila culturing vials medium is most sensitive to user manipulation. Excessive force should be avoided.

Troubleshooting:

Possible problems, causes, and solutions are presented in Table 1.

Table 1.

Troubleshooting Guide for Colorimetric Synchronization of Drosophila Larvae

Problem	Possible Cause	Solution
Low embryo yield	Older adults used in cage; suboptimal quantity of adults chambered within cage	Fecundity declines with age. Freshly eclosed adults should be used. Titrate numbers of adult Drosophila within cage to optimize embryo yield
Low first instar larvae yield	Developmental delay onset by genetic background; unstable environmental conditions	Conduct experiment in parallel replicates and/or increase number of Drosophila within cage to increase embryo yield; clean up older strains by backcrossing; ensure proper functioning of incubator; add a humidity source as needed
Low second- third instar larval yield	Physical manipulation during transfer of first larval instar to culturing vial; contaminated yeast paste	Minimize manipulation of first instar larvae during transfer; use gloves and clean tools when handling yeast paste

Understanding Results:

Please see information about sample data in the Understanding Results section (below).

Volumetric measurement of the third instar central nervous system

Drosophila neurogenesis occurs through three distinct developmental waves commencing during embryogenesis, when neural stem cells (NSCs) and glia are born (Homem, Repic, & Knoblich, 2015; Landrum et al., 2016). A second neurogenic wave during the first larval instar augments the number of NSCs within the developing larvae (Kohwi & Doe, 2013; Li & Hidalgo, 2020). During the third and final neurogenic wave, NSCs stop dividing (del Valle Rodriguez, Didiano, & Desplan, 2011). The balance between self-renewal versus differentiation is governed by the segregation of cell fate determinants along the apical-basal polarity axis and is vital to maintain homeostasis in neurogenesis, where deregulation can lead to tumorigenesis, neurodegeneration, or neurodevelopmental disorders, such as microcephaly (Cabernard & Doe, 2009).

Drosophila have been used to study the neurodevelopmental mechanisms of human diseases, including microcephaly (Link & Bellen, 2020). Here, we show our protocol can be employed to assay microcephaly.

For this study, age-matched larval brains were identified by partial gut clearance, dissected, fixed, and stained for DAPI as a marker for brain volume (Lerit, Plevock, & Rusan, 2014; Link et al., 2019). Entire brain volumes were then imaged on a spinning disk confocal microscope. Images from 30 brains per genotype from control and mutant lines were randomized and brain volumes measured in the Imaris image analysis package using the Surfaces tool. Statistical variance was assessed by t-test in GraphPad Prism software with p<0.05.

Our protocol permitted the comparison of brain volumes from control versus mutant samples in which a null mutation in the orb2 gene resulted in a significant (24–36 hour) developmental delay. Age-matched control versus orb2 mutants show significantly different third instar larval brain volumes (Figure 3; (Robinson et al., 2022)).

Figure 3. orb2-dependent microcephaly in larval brains.

(A-B) Images show control and orb2 null third instar larval brains stained with DAPI. The optic lobe is outlined (dashed line). Bars: 40 μm (C) Graph shows quantification of brain volumes from age-matched (partial gut clearance) samples, where each dot represents a measurement of a single optic lobe from N=30 brains per genotype normalized to the wild-type (WT) control. Loss of orb2 results in significantly smaller brain volumes by t-test analysis. Data shown are adapted from (Robinson et al., 2022).

Exploring Ecdysone Receptor Activity in third instar wing imaginal discs

A combination of systemic and local signals coordinates the growth and patterning of tissues during development. The steroid hormone 20-hydroxyecdysone (20E) is the central regulator of developmental transitions in Drosophila (Riddiford, 1993). 20E binds the Drosophila nuclear hormone receptor Ecdysone Receptor (EcR) to activate the expression of target genes (Koelle et al., 1991). In the absence of hormone, EcR is bound by transcriptional repressors to repress transcription (Tsai, Kao, Yao, McKeown, & Evans, 1999); (Schubiger, Carré, Antoniewski, & Truman, 2005). 20E-EcR signaling elicits a wide range of tissue-specific responses during larval development in a stage- and tissue-specific manner. In imaginal wing discs, bilaminar sheets of epithelial cells which are precursors to the adult wing, 20E-EcR signaling drives proliferation, morphogenesis, and differentiation during third instar development (D’Avino & Thummel, 2000; Herboso et al., 2015).

The importance of 20E-EcR signaling in developmental timing has made examining the molecular mechanisms of 20E-EcR signaling challenging. For example, EcR loss-of-function alleles or inhibition of 20E production leads to developmental arrest, which confounds tissue-specific studies of their molecular roles. To analyze 20E-EcR roles in developing tissues, we created a transgenic tool that allows for tissue-specific expression of a fragment of EcR containing the ligand binding domain and sequences predicted to interact with coactivators and corepressors (EcR^LBD). We used this tool to reveal an activation-to-repression switch that occurs during a relatively short temporal window within the third instar larval period (Wardwell-Ozgo et al., 2022).

Our colorimetric synchronization protocol was key to comparing responses of EcR target genes via the EcR^LBD probe. Briefly, age-matched control (RFP) versus experimental (EcR^LBD) expressing third instar larvae were identified by food clearance in the gut, and wing discs were dissected, fixed, and immunostained with anti-β-galactosidase antibodies to detect activity of the EcR reporter, EcRE-lacZ. Optical (z-stack) projections were collected on a confocal microscope. Use of the colorimetric synchronization protocol was thus key to demonstrating significant alterations in 20E-EcR activity in larval wing cells expressing EcR^LBD (Figure 4; (Wardwell-Ozgo et al., 2022)).

Figure 4. EcR^LBD reveals an EcR repression-to-activation switch in third instar larval wing discs.

Optical (z-stack) projections of third instar larval wing discs immunostained with anti-β-galactosidase (LacZ; green) to detect EcRE-lacZ expression in control (en>RFP; A-C) or en>EcR^LBD (D-F) discs. RFP reporters are displayed in red and mark the posterior compartment. Images are representative from N=16 control dark, 19 partial, and 7 clear versus N=6 EcR^LBD dark, 15 partial, and 7 clear third instar larvae. Note how in the control samples, LacZ expression increases over time. In contrast, expression of EcR^LBD results in posterior enrichment of LacZ in early (dark) larvae, while anterior enrichment is observed during middle and late (partial and clear, respectively) third larval instar development. Data shown are adapted from (Wardwell-Ozgo et al., 2022).

[*Copyeditor: Please double check with authors regarding the permission to use the adapted figures 3 and 4. I asked them and here is their response: “These figures contain data uploaded to Biorxiv by authors of this protocol, accessible under a CC-BYNC 4.0 license.” If this is the case, it should be mentioned in the Acknowledgements section and/or in the legends]

Time Considerations:

This protocol can be completed over the course of 7 days. On day 0, adult Drosophila of genotypes of interest are housed within embryo collection cages. On day 1, the cage is fed, and the flies acclimatize to their conditions. On day 2, timed embryo collections commence. By day 3, first instar larva are transferred to culturing vials containing blue yeast paste and allowed to develop at 25°C. During days 3–4, larvae complete the first and second instar phases such that, by day 5, larvae enter the third instar stage and may be selected for analysis. By the end of day 7 larvae complete the third instar stage and enter pupariation. The median time to pupariation is 120 hours AEL under standard conditions.

DATA AVAILABILITY STATEMENT:

The data that support the protocol are openly available on Biorxiv at https://doi.org/10.1101/2021.11.23.469707 and https://doi.org/10.1101/2022.04.07.487542.