Arthropod Lab Part1: Flies
Nipam Patel [email protected] Matthew Ronshaugen [email protected]
Justin Bosch [email protected] Katherine Brown [email protected] Heather Bruce [email protected] Erin Jarvis [email protected] Ryan Null [email protected] TABLE OF CONTENTS
I. INTRODUCTION 2 II. SCHEDULE 3 III. EXPERIMENTAL OVERVIEW 4 IV. PROTOCOLS 9 IV.1 General Fixation and Antibody Staining 9 IV.2 Rapid Antibody Staining Protocol 11 IV.3 Fixing and Staining Post-embryonic Tissues (Imaginal Discs) 12 IV.4 In situ Hybridization 14 IV.5 EdU labeling of cell proliferation in tissues 16 V. GENERAL SOLUTIONS 17 V.1 Fixatives 17 V.2 Solutions for antibody protocols 17 V.3 Solutions for in situ hybridization 19 VI. MAKING DISSECTIONS TOOLS 20 VI.1 Making blunt probes 20 VI.2 Making Tungsten needles for dissecting 20 VII. AVAILABLE STOCKS & REAGENTS 21 VII.1 Antibodies Separate Handout VII.2 In situ Probes 21 VII.3 Fixed Embryos 26 VII.4 Drosophila Stocks 27 VIII. DROSOPHILA DEVELOPMENT & STAGING 28 VIII.1 Embryogenesis 28 VIII.2 Larval Development and Morphogenesis 32 VIII.3 Adult Morphology 34 VIII.4 Female Reproductive System 35 VIII.5 Male Reproductive System 37 VIII.6 Imaginal Disks and Generation of Mosaic Tissues 38
1 I. Introduction In this module, you will learn about a variety of arthropod systems, including the model genetic system, Drosophila melanogaster. Most importantly, we would like you to leave with the ability to analyze and compare the development of different arthropod embryos. In order to do that, you will be performing various molecular and embryological techniques, such as antibody staining, in situ hybridization, live imaging, and lineage tracing using both classical injection and modern genetic methods. At first we would like you to use a set of antibodies to detect the expression of important developmental proteins and RNAs in the fruit fly Drosophila melanogaster. This will allow you to master the procedure of antibody staining, examine the spatiotemporal expression of these proteins, and study the development of various tissues. Because many of these proteins are strongly conserved in all arthropods studied to date, a subset of these antibodies allow you to go crazy and stain all sorts of arthropods that cross your way!! We will have many critters from which you can collect embryos and make them shine! In this module you will also have the opportunity to look at aspects of post-embryonic development. In particular, we will look at wing imaginal disc development in Drosophila and scale patterning in butterflies. You can stain imaginal disks with available antibodies and compare the expression pattern of various proteins between flies and butterflies, and look at butterflies scales by scanning electron microscopy. We will also give you several ideas for “projects” that we can discuss on the evening of the first day of the lab, but we also encourage you to try out as many techniques as possible and to look at as many critters as you can. We have provided an extensive list of possible experiments in section III. We suggest you read through them all and see what captures your interest. Do not try to do too much; by no means do we think you can or should do all of them. So be selective to optimize your time here. Talk to use about your goals, and we will help you plan and execute your experiments.
2 II. Schedule Monday (lecture: Chris Rushlow) Begin rapid staining Look at pre-stained Drosophila embryos (demo of mounting) Review of available antibodies Complete and begin imaging rapid stains Dissection of Drosophila imaginal disks (demo at 8 PM) Set up overnight staining, live imaging, and embryo collections
Tuesday (lecture: Alan Spradling) Continue antibody staining Introduction to in situs Volocity training (2:30 – 4:00) Introduction to fly genetics and crosses Parhyale and Mysid dissection (demo at 5 PM) Ovary dissection (demo at 8 PM)
Wednesday (lecture: Iswar Hariharan)
Arthropod diversity (inc. plankton tow; demos at 8 PM) Parhyale injections (demos at 2PM) Dissection of butterfly imaginal disks (demo at 4 PM)
Thursday (lecture: Nipam Patel)
Continue experiments Softball practice or snorkeling (weather permitting)
Friday (lecture: Matt Ronshaugen) Continue experiments
Saturday (lecture: Nipam Patel) Continue experiments Show N’ Tell
3 III. Experimental Overview
III.1 Observations of Embryogenesis A. RAPID Antibody Staining of Drosophila Embryos. In this experiment, you will investigate protein expression patterns throughout Drosophila development that illustrate the following gene classes/organs (appropriate antibody in parentheses): maternal gradient (Rb anti-Bcd), pair-rule (Rb anti- Ftz, Rb anti-Eve), segment polarity (M anti-wingless [4D4]), Hox (M anti-Antp [8C11], Rb anti-teashirt), CNS (anti-HRP), and muscle (Ratb anti-tropomyosin). During the Arthropod Module, you will see more examples of these patterns in Drosophila and examine whether other arthropods share them as well.
To do the experiment, split into six groups of four. Each group should complete all four of the assigned antibody stains on Drosophila, plus one control tube (no primary antibody). The rapid antibody protocol is in section IV of this manual, but is repeated below. Use the primary antibodies at the "60 min" dilutions listed on the antibody table. Use secondaries at 1:500 (as opposed to the standard 1:1000). NOTE: This rapid protocol only works for a few antibodies. Most every antibody will work better using the standard protocol (overnight in primary, 2 hrs in secondary).
The fly embryos you will be given are a mix of embryos ranging from 0-18 hours and are in methanol. The tube contains enough embryos for the five stains you will do here. When carrying out fly embryo staining in eppendorf tubes, plan on having 15µl of settled embryos in MeOH per 1.5ml eppendorf tube. This will be about 20µl when rehydrated. Using too many or too few embryos can affect the signal-to- noise ratio of your staining.
1. Rehydrate 3X 1 min followed by 1X 10 min with PT.
2. Incubate 10 min in 200 µl PT+5% NGS.
3. Add primary antibody to the appropriate final concentration. ("60 min" concentration)
4. Mix and incubate in the primary antibody at room temperature for 60 min.
5. Wash 3X 1 min with PT.
6. Wash 3X 10 min with PT.
7. Add secondary antibody (in 200 µl of PT+ 5% NGS). It is not necessary to pre-block with PT+NGS. Use fluorescently labeled secondaries at a dilution of 1:500 when using this rapid protocol (as opposed to 1:1000 with the "normal" protocol).
8. Wash 3X 1 min with PT.
9. Wash 3X 10 min with PT.
10. Add 200 µl 50% glycerol with 1.0 µg/ml DAPI and incubate 15 min.
11. Remove 50% glycerol and add 70% glycerol (no DAPI). Embryo will be ready to view in about 15 minutes.
4 B. Molecular Markers of Embryonic Development
Using a specific set of antibodies you will be able to see the expression of proteins involved in different steps of early development, such as the formation of the body segments, the specification of the different body regions (read, thorax and abdomen) as well as the formation of neurons and axons. We provide you with a list of additional antibodies for you to expand your expression analysis. You can either select a few genes and compare them across many different species or, pick a few arthropod species and study the expression of many genes.
Gap and Pair Rule Look at the expression patterns of gap and pair-rule genes during early Drosophila development. Look at the expression of gap and pair-rule orthologs in other arthropods (you are encouraged to look in other phyla as well).
Pair Rule and Segment Polarity Examine the expression patterns of Pax 3/7 and Even-skipped orthologs in a variety of organisms. How does this compare with the expression of Engrailed orthologs across species? What does this say about the evolution of segmentation in arthropods?
Hox and Appendage Formation Examine the expression of Hox genes in Drosophila and other arthropods. Examine how Hox gene patterns have changed during evolution and the possible morphological consequences of these changes.
Neurogenesis and Axonogensis Examine the process of Drosophila neurogenesis and axonogenesis. Compare neurogenesis and axonogenesis in Drosophila to neurogenesis and axonogenesis in other arthropods
C. Live Imaging of Drosophila Embryos Collect Drosophila embryos at the appropriate stage (as close to fertilization as possible if you want to look at early nuclear divisions; late blastoderm if you want to watch gastrulation, mid germband extension for tracheal development, early germband shortened to view dorsal closure), mount in petri dishes with glass bottom coverslips coated with heptane glue, fill with Schneider's insect media (room temp) and image away on confocal or inverted widefield microscope. Try live imaging the following: 1) Wild type 2) GFP and RFP lines (e.g. moesin:GFP, histone:RFP, hh-GFP, Btl-GFP is particularly excellent) 3) GAL4 lines (e.g. Dpp-Gal4, hh-Gal4) crossed with UAS-RFP or G-TRACE.
D. GAL4 drivers in development, in live or fixed animals The GAL4/UAS binary system has been adapted from yeast for use in flies. Expression of the GAL4 gene is driven by a specific promoter (e.g. hsp70, tubulin, actin, nubbin, etc.). GAL4 binds to Upstream Activating Sequences (UAS) and activates transcription of genes adjacent to theses sites. This allows for any gene placed downstream of UAS to be expressed in a specific tissue or at a specific time point in development. In Drosophila a wide range of different GAL4 drivers and UAS transgenes are available. 5 Suggestions: 1) Express GFP or RFP in specific tissues by crossing GAL4 and UAS-GFP lines 2) Express Ubx ectopically in specific tissues by crossing GAL4 and UAS lines 3) Observe live during development (look at GFP), or fix and stain with interesting antibodies to observe GFP plus expression of another gene The GAL4/UAS system has been combined with the FLP/FRT system for producing mitotic recombination (see VIII.2 for a description) to produce powerful lineage tracing tools. By driving the expression of FLP with GAL4, you can permanently label a given cell and all its progeny with GFP. Using “G-TRACE” you can permanently mark cells that have expressed a given GAL4 and simultaneously view whether these cells or others are currently expressing this GAL4.
E. Analysis of mutant embryos. How is development disrupted by the lack of early patterning genes? We have fly stocks that carry mutations in various early embryonic patterning genes. You can collect homozygous mutant embryos and perform combinations of cuticle preparations, antibody staining, in situ hybridization, live imaging, and anything else you can think of, to analyze their phenotypes! For a full list see section VII.4. Available to you will be a series of segmentation mutants as well as several effecting neural development.
F. in situ Hybridization Assay gene expression for any number of genes in wild type and mutant embryos. You can visualize both cytoplasmic transcripts (exonic probes) and nascent transcripts (intronic probes). You will be provided with pre-fixed embryos in methanol, but you can also collect your own from the stocks available if you prefer, or for custom combinations of your choice. The protocol provided is suitable for single, double, or triple label, or a combination of antibodies and in situ probes. You should double check the fluorophores available (see Antibody list) for compatibility with the lasers and filter cubes on the microscopes. Consult with Matt to check if your fluorophore combination is possible. Keep in mind that this is a three-day protocol requiring two overnight steps, so plan accordingly! Start thinking about your probe combinations on MONDAY and start the first step (hyb overnight) on TUESDAY. A list of all the genotypes and probes available is in the back of the manual. For many genes we have probes for flies, beetles, and bees. Why not see how the expression patterns have evolved, and look at your gene of interest in different insect orders? Evo-Devo fun! Suggested Experiments: 1) How about a multiplex in situ to assay what happens to gastrulation in segmentation mutants? 2) Various developmental processes: e.g. we have a number of probes and antibodies to look at heart formation (tinman) pericardial cells (eve) and muscles (mef2, twist) across the various species 3) Try looking at the transcription of the 5’ and 3’ end of the same gene. This is particularly interesting for long genes (Antp 105KB and Ubx 84KB) in early embryos when cell cycles are short. 4) Observe the border of the mesoderm and neuroectoderm by visualizing rho, sim and snail (or twist). Also you can examine what happens when some of the early mesodermal DV transcription factors expressed in an AP gradient. 5) Observe the differential expression of the related but differentially expressed genes SoxN and Dichaete in the nervous system. 6) Compare the expression pattern of midline patterning genes (e.g. sim) between Drosophila and Apis embryos.
6 G. Wing Imaginal Disc Staining Look at the expression of Engrailed and Ubx orthologs via antibody staining in Drosophila and butterflies. Engrailed marks the posterior wing compartment (and, in butterflies, the center of the eyespot), and Ubx is expressed in the T3 wing/haltere, but not in the T2 wing. Stains to do and compare: 1) 4F11 (anti-Engrailed) on Drosophila wing discs (3rd instar) 2) 4F11 (anti-Engrailed) on butterfly wing discs (4th and 5th instars) 3) FP6.87 (anti-Ubx) on Drosophila wing discs (3rd instar) 4) FP6.87 (anti-Ubx) on butterfly wing discs (4th and 5th instars) Notes: You can stain Drosophila and butterfly discs in the same tube Put fly embryos in as controls (will stain for Engrailed) Only late 5th instars and older butterfly larvae will have eyespot staining. For best eyespot stains, use discs from 12hrs post pupariation.
H. Imaging multi-color clones in Drosophila imaginal discs. See Section VIII.6 for a description of how clones are generated in imaginal discs.
We brought a multi-color genetic labeling system called “LOLLIbow” (a Drosophila variant of Brainbow) for you to experiment with, see section VIII.6. This allows you to simultaneously image (live or fixed) multiple clones in tissues with three different fluorescent proteins (mCerulean, EYFP, and mCherry). We have three versions of LOLLIbow - in the wing imaginal disc (rotund-Gal4), the hedgehog expressing domain (hedgehog-Gal4), and all neurons (elav-Gal4). We will supply 3rd instar larvae and pupae with multi-color clones in the wing imaginal discs (we will induce clones in 48hr old embryos), but consider looking in different tissues or at different developmental timepoints (e.g. embryos). Here are a few suggested experiments.
1) Live image multi-color clones in the pupal wing disc. Boulina et al. 2013 show wing imaginal disc cells extending filopodia towards each other and extracellular puncate (exosomes?) from differently labeled clones. Try to replicate this experiment as follows: Find a pupae (12-24hrs after pupa formation) expressing OFP. Carefully dislodge it with a metal probe. Wash pupae with PBS and mount in a glass- bottomed 25 mm Petri dish with a layer of thiodiethylene glycol. Place a coverslip on the sample and image for X hours.
2) Observe multiple clones in fixed wing imaginal disc (using rotund-Gal4) and stain with anti-Ci to mark the compartment boundary. Do any clones lie right on the compartment boundary? Is there a pattern to the shape/size/orientation of the clone for where the clone lies in the wing pouch?
3) Fix and mount larval or adult brains and try to find image multi-color neurons.
7 J. Mosaic analysis of growth regulators in the eye imaginal disc
We brought a genetic system that automatically generates mosaic tissue expressing GFP and a gene of interest in the eye imaginal disc, called CoinFLP (see Section VIII.6). One dramatic application of this technique is to overexpress positive regulators of growth, producing overgrowth of the GFP tissue compared to the neighboring wildtype non-GFP tissue. Overgrowing cells can sometimes have a competitive advantage over neighboring wildtype cells (for some, but not all, growth regulators), leading to their elimination by apoptosis.
Dissect eye imaginal discs from 3rd instar larvae resulting from different crosses of CoinFLP UAS-GFP and UAS-X lines. Overgrowing cells are marked by GFP. Use markers of proliferation to see if cells are more frequently in S-phase (anti-phospho-Histone H3), and markers of apoptosis to see if overgrowing or wildtype cells are undergoing cell death (anti-cleaved Dcp1). Feel free to try other antibody stains as well.
8 IV. Protocols
IV.1 General Antibody Staining Fixation: Not fixing the embryo sufficiently will result in high background levels and over fixation may prevent your embryos from staining at all – so find a time range that works for you and also be consistent when you start timing (i.e., when most of the embryo is exposed to fix). After fixing, most washes contain detergent. This helps to prevent fixed embryos from sticking to pipettes. You can introduce detergent in the last 2-5 minutes of fixation by adding 1/5 volume of PT to your fixative. 1. Transfer embryos from agar collection plates into a nylon mesh basket using water and a small paintbrush. 2. Place the egg baskets in a small glass beaker partially filled with 50% bleach solution. Gently swirl the basket or use a Pasteur pipette to rinse the embryos. Dechorionation should take about 3 min; however, the potency of bleach varies so monitor the process under a dissection scope and stop it once the chorion has dissolved away. 3. Immediately wash thoroughly with room temperature water. 4. Transfer the embryo to a 20 ml glass scintillation vial containing 10 ml of heptane and 10 ml of PEM- FA (fixative solution, see section V). You can do this by shaking the mesh directly into the heptane phase and can use heptane to wash down any embryos stuck to the side of the basket. 5. Mix gently for 15-20 minutes. 6. Remove the aqueous phase (lower phase). Add more heptane if needed to maintain a volume of at least 8 mls of heptane. Try to remove all of the aqueous phase. 7. Add 10 ml of methanol and shake vigorously for 15-30 seconds. Devitellinized embryos will fall to the bottom (methanol phase). 8. Pipette out the embryos from the bottom and transfer them to a new tube. 9. Wash embryos several times with methanol to remove traces of heptane. 10. Embryos can be stored in methanol at -20°C for several years. Rehydration and Staining: 1. Rehydrate embryos from methanol with 3 X 5 minute PT washes. Only rehydrate what you need for today, leaving rest in methanol for future use. As a rule of thumb, 15 µl of settled fly embryos in MeOH will be about 20 µl when rehydrated, and this 20 µl volume is what you want per 1.5 ml eppendorf microcentrifuge tube. If working with an arthropod other than Drosophila, add some fly embryos (10-30 embryos) to the tube as well (they will act as an internal control.) 2. Incubate 10-30 min in 300 µl of PT+N (PT + 5% NGS). The normal goat serum (NGS) will help to block nonspecific antibody binding sites. Gently mix by spinning the tubes. Avoid shaking or flicking the tubes as the embryos will splash up onto the walls of the tube and dry out resulting in either unstained or non-specifically stained embryos. 3. Add the appropriate amount of primary antibody to achieve the desired final concentration (see antibody table). 4. Gently mix the embryos and antibody solution and incubate overnight at 4°C. 5. Wash 3 X 1 min with PT. Before these washes are started, it is possible to recover the diluted primary antibody, and this used antibody can often be re-used several more times. Store this diluted antibody at 4°C. 9 6. Wash 3 X 30 min with PT. 7. Incubate 10-30 min in 300 µl of PT+NGS as in step 2 above. 8. Add appropriate secondary antibody to the proper final concentration (1:500 - 1:1000 for most of the Alexa conjugated secondaries). 9. Mix the embryos and secondary antibody solution gently and incubate for 2 hrs at room temperature. 10. Wash 3 X 1 min with PT. 11. Wash 3 X 30 min with PT. 12. If you used a fluorescently tagged secondary antibody, add 200 µl 50% glycerol with DAPI (either 0.1 or 1.0 µg/ml DAPI) for 15-20 minutes, and then replace with 300 µl 70% glycerol (no DAPI). AlexaFlour conjugates are very fade resistant even without the addition of an anti-fade compound.)
10 IV.2 Rapid Antibody Staining Protocol
While this protocol produces antibody stains in just a few hours, it only works well on very robust antibodies. Use this only for antibodies listed with a "60 min" concentration on the antibody list.
12. Rehydrate with 3X 1 min followed by 1X 10 min with PT.
13. Incubate 10 min in 200 µl PT+5% NGS.
14. Add primary antibody to the appropriate final concentration. ("60 min" concentration)
15. Mix and incubate in the primary antibody at room temperature for 60 min.
16. Wash 3X 1 min with PT.
17. Wash 3X 10 min with PT.
18. Add secondary antibody (in 200 µl of PT+ 5% NGS). It is not necessary to pre-block with PT+NGS. Use fluorescently labeled secondaries at a dilution of 1:500 when using this rapid protocol (as opposed to 1:1000 with the "normal" protocol).
19. Wash 3X 1 min with PT.
20. Wash 3X 10 min with PT.
21. Add 200 µl 50% glycerol with 1 µg/ml DAPI and incubate 15 min.
22. Remove 50% glycerol and add 70% glycerol (no DAPI). Embryo will be ready to view in about 15 minutes.
11 IV.3 Fixing and Staining Post-embryonic Tissues (Imaginal Discs)
A. Drosophila Wing Imaginal Discs
Dissecting wing discs: On a Slygard coated dish in 1X PBS, dissect imaginal discs by one of the following methods: Method A: • Tear larvae in half (cut across the “waist”) with forceps • Turn the anterior half of the larva “inside-out” by pushing in at the head/mouth-hooks with the tip of one pair of forceps while drawing the cuticle back over this pair of forceps with a second pair.
• Gently remove the fat body, salivary glands and gut, leaving the brain and imaginal discs intact. These are at the anterior end of the larva near the mouth hooks. (Tip: leave the main lateral trachea in place as this will almost always guarantee recovery of the wing discs) • The wing discs are pinned to the sides of the larva by the two prominent lateral trachea. They are the largest imaginal discs. • The eye-antennal discs are attached to the optic lobes of the brain and the mouth hooks. Method B: • Grab the mouth hooks with one pair forceps • Grab in the middle of the larval body with 2nd pair forceps • Pull mouth hooks (and connected imaginal discs) out of body while holding the body in place with your 2nd pair of forceps. • Wing discs will be in a bunch with other discs and brain. Clean away any extra tissue such as fat body or salivary gland. The wing disc is largest and has a prominent pocket. • This method is preferred for dissecting eye-antennal discs. Ideally your discs will spend no more than 15-20 min in PBS before you begin your fixation. The importance of this will vary depending on the antibody you are looking at. You can either do the initial dissection (inverting carcasses or pulling mouth hooks), then add fix and clean off the extraneous tissue while fixing or you can complete your dissections in PBS and transfer just the carcass + discs or mouth hooks + discs into an eppendorf for fixation. How you do this will depend largely on personal preference and dissection speed/skill.
12 Fixation and Staining: 1. Fix wing discs or eye discs (either attached to the inverted carcass for A or to the mouth hooks and other discs for B) in 4% PFA in 1X PBS for 15-20 minutes. 2. Wash at least 3X 10 min in PT to remove residual fix. 3. Block for 30 min in 10% NGS in PT. 4. Add the appropriate amount of primary antibody to achieve the desired final concentration, mix gently and incubate overnight at 4°C. 5. Rinse once in PT, then wash 3X 10 min in PT. 6. Block for 30 min in 10% NGS in PT. 7. Add the appropriate secondary antibody to the proper final concentration, mix gently and incubate with for 2hrs at room temperature or overnight at 4°C. 8. Rinse once in PT, then wash 3X 10 min in PT. 9. Incubate in 50% glycerol (0.1 or 1.0 µg/ml) with DAPI for 15-30 minutes 10. Mount in 70% glycerol. Transfer carcasses/mouth hooks to slide in glycerol and dissect off the desired imaginal discs. If the 3-dimensional morphology of the disc is important use spacers (either double- sided sticky tape or two coverslips) to hold coverslip.
13 IV.4 In situ hybridization Day 1: (3 hours) 1. Wash 1 time in 1:1 MeOH:EtOH for 5 minutes 2. Wash 1 time in EtOH for 5 minutes 3. Wash 1 time 9:1 Xylenes:EtOH for ~ 1 hour 4. Wash 2 times in EtOH for 5 minutes 5. Wash 1 time in 1:1 MeOH:EtOH for 5 minutes 6. Wash 1 time in MeOH for 5 minutes 7. Wash 1 time in 1:1 Fix Solution:MeOH for 5 minutes 8. Fix for 25 minutes in Fix Solution 9. Wash 3 times in PT 10. Wash 2 times in 1:1 PT:Hyb Buffer for 5 minutes 11. Wash 2 times in Hyb Buffer at 55 degrees 12. Wash 1 time in Hyb Buffer at 55 degrees for 30 minutes 13. Incubate embryos in Probe Solution at 55 degrees for at least 16 hours (overnight) Day 2: (6 hours) 14. Wash 5 times in Hyb Buffer at 55 degrees for 1 hour total (~4 15 minute washes) 15. Wash 2 times in 1:1 PT:Hyb Buffer for 5 minutes (now at room temp) 16. Wash 3 times in PT 10 minutes each 17. Wash 3 times in PT+WBR for 20 minutes (1 hour total) 18. Incubate in Antibody Solution at 4 degrees for 12-18 hours (overnight) Day 3:Multiplex Fluorescent detection 19. Wash 3 times in PT 10 minutes each 20. Wash 3 times in PT+WBR for 20 minutes (1 hour total) 21. Incubate in secondary Antibody Solution at room temp for 2-3 hours. 22. Wash 3 times in PT 23. Wash 2 times in PT 20 minutes each 24. Mount in Prolong Gold w/DAPI
General Hints and Notes on in situ hybridization experiments • The fix to use is 1:1 PBS:10% Ultrapure Formaldahyde • Combine in situ with antibody staining to detect both RNA and protein • in situ hybridizations (ISH) and ISH/Ab combos need to be started on the 2nd afternoon (Tuesday). If you are doing Ab stainings only, you can start those as late as the 3rd day (except for the group stains that are the introduction to the Fly and Arthropod Modules to be done on Monday). 1. The first day consists of mostly washes, so you should be able to continue washes during the evening labs and subsequent days if you need to. 2. Unless otherwise indicated, washes are not all that time-sensitive, especially when they are pure alcohol or PT washes. However, do try to remain in the ballpark of the times indicated. • Choose your probes and get started. • Map out your desired staining protocol ASAP so you can identify any potential problems
a. You are entirely free to do design your own experiments and do as many samples as you like. However, as you will be dealing with different methods/experimental setups/protocols simultaneously, we suggest doing 4 to 8 samples. Try to do at least one colorimetric ISH, one fluorescent ISH, and one Ab stain/ISH combo. b. Do (or get one of your friends to do it and show you) a WT (i.e. yw) control in parallel with mutant genotypes. 14 c. You may want to compare protein and RNA expression patterns. • Remember that we also have cytoskeletal stains and nuclear stains such as DAPI. Some of the mounting media (VectaShield) contain DAPI. An additional way to visualize nuclei is to stain with the anti-lamin antibody. • You$can and should make use of the online resources to get a better idea/more information about the genes you are dealing with... start with www.flybase.org. 26 Multiplex in situ Hybridization: These are just some of the available options/combinations that should work. Please feel free to make up your own staining schemes, and ask a TA if you have any questions.
Hapten 1o Antibody 2o Antibody Alexa Cojugate 3 Probe Labels DIG Sh 〈 DIG Dk 〈 Sh 555 BIO Mo 〈 BIO Dk 〈 Mo 488 DNP Rb 〈 DNP Dk 〈 Rb 647 FITC Rb 〈 DNP Dk 〈 Rb 647 Mo = Mouse; Rb = Rabbit, Sh = Sheep; Dk = Donkey
This is the basic triple. All primaries and secondaries should be diluted 1:400. To do a single or a double, simply eliminate the primary and/or secondary antibodies for that channel. A general rule given these combinations: • DIG will be the strongest/cleanest • DNP will be good but not visible to the eye (when detected with Alexa 647) • BIO (Biotin) and FITC (Flourescien) can be weak/backgroundy occasionally (late embryos have significant background in the yellow/green range) To include an antibody in your in situ, it is often best to substitute it in place of the Mouse anti- BIO antibody. This is because many antibodies are mouse monoclonal and the BIO channel will detect mouse primaries. Other combinations of primary and secondary are possible, but those above tend to work consistently.
15 General Solutions V.1 Fixatives Formaldehyde Fixes: 9:1 PBS fix = 9 parts 1X PBS: 1 part 32% formaldehyde OR 9:1 seawater fix* = 9 parts filtered seawater: 1 part 32% formaldehyde
Formaldehyde = 32% Fisher cat # F79-500 *saltier fixes are for Parhyale and other marine creatures
10X PBS: 18.6 mM NaH2PO4 (2.56 g NaH2PO4 . H2O per 1000 ml dH2O) 84.1 mM Na2HPO4 (11.94 g Na2HPO4 per 1000 ml dH2O) 1750.0 mM NaCl (102.2 g NaCl per 1000 ml dH2O) Adjust pH to 7.4 with NaOH or HCl. Prepare 1X PBS by diluting 1:10 with dH20. Both 1X and 10X PBS can be kept indefinitely at room temp.
PT: 1X PBS 0.1% Triton X-100 Mix 100 ml 10X PBS, 899 ml dH2O, and 1 ml Triton X-100. Store at 4°C or at room temp.
PT + 5%NGS (PT+N): 1X PBS 0.1% Triton X-100 5.0% Normal Goat Serum (Gibco-BRL Cat. No. 200-6210AG) Heat inactivate the serum at 56°C for 30 min. Filter through a 0.22 µm filter while still warm. Aliquot into sterile tubes. Store aliquots at –20°C. Once thawed, aliquots are stable for several months at 4°C. To prepare the PT+NGS solution, mix 4.75 ml PT with 0.25 ml Normal Goat Serum and store at 4°C. Solution will usually last at least two or three weeks. Discard if bacterial growth is detected (solution will turn cloudy).
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16 Glycerol solutions: Some batches of glycerol contain contaminants that cause nickel-enhanced DAB reactions to fade within a day or two. To avoid this, use ultrapure glycerol (Boehringer Mannheim, Cat. No. 100 647). Prepare 50% and 70% glycerol solutions by mixing the appropriate volumes of glycerol with 1X PBS. Use pH paper to make certain that the pH of the glycerol solutions is around 7.4. Low pH will cause rapid fading of DAB reaction products. Glycerol solutions can be stored at room temperature. Glycerol solutions with DAPI should be stored in dark at 4°C.
Glycine pH 2.0: 0.375 g Glycine 250 µl 20% Tween-20
Dissolve Glycine in 40 mL dH2O and adjust pH to 2.0 with concentrated HCl. Add Tween-20 and adjust volume to 50mL with dH2O
17 V.3 Solutions for in situ hybridization
Fix Solution (10mL): PT 9ml 32% Formaldehyde 1ml
20X SSC (50mL): NaCl 8.77g Sodium Citrate 4.41g MilliQ Water fill to 50mL
20X SSC pH=4.5 (14mL): (make fresh) 20X SSC 14mL Concentrated HCl 225uL
Hyb Buffer (40mL): Formamide 20mL 20x SSC (pH 4.5) 10mL 10% SDS 400uL milliQ H2O fill to 40mL
Probe Solution (500ul [1:500]): RNA probe 1ul Hyb Buffer 500ul adjust concentration accordingly
Antibody Solution (3mL): anti-DIG antibody 1ul
18 VI. Making Dissection Tools
VI.1 Making blunt probes 1. Heat a long glass pipette with a Bunsen burner and pull so that the long skinny part stretches and breaks. 2. Now there are two pieces of what used to be a long glass pipette. 3. With the larger stem piece, heat the newly formed end so that it rounds up. The objective here is to get a small round end so the animals don’t get hurt.
VI.2 Making Tungsten needles for dissecting (station in back of the main lab)
Electro-chemically sharpened: 1. Use Tungsten wire 0.005” diameter (Ted Pella, Inc., the Electron Microscopy Supply Center; Redding, CA. 1 2. Thread wire through a 26 G x /2 Needle (from Precision Guide #305111). Crook the back end of the wire so that it stays inside. Attach to a 1 cc syringe. 3. Set up a beaker with 1 N NaOH. WEAR SAFETY GOGGLES TO PREVENT SPLASHING OF NaOH INTO YOUR EYES! 4. Hook electrical clamps: one clamps to the beaker and touches NaOH solution; the other is clamped to the needle. BE CAREFUL NOT TO TOUCH THE TWO CLAMPS TOGETHER OR YOU ARE IN FOR A SHOCK!!! (you will also short out the transformer) 5. Plug clamps into the Variable Auto Transformer. Set input = 120 V, 50/60 Hz; Output = 0-120/140 V, 10 A, 1.4 KVA. 6. Put switch on 120 V. Set dial to at least “2”, but no higher than 6. 7. Dip needle into NaOH – where you see bubbles is where the metal is dissolving. A steady ‘up and down’ motion will ensure that the tip of the needle is the sharpest.
19 VII. Available Reagents
VII.1 Antibodies (see separate PDF)
VII.2 In situ Probes
LEGEND FOR NUMBERED PROBES PROBE LABEL Gene DNP DIG Biotin Fitc 1 krupple intron Y Y 2 knirps intron Y Y Hunchback 3 intron Y 4 Hunchback 3' Y Y 5 Antp 3' Y Y(WEAK) 6 Antp 5' Y Y Y Y 7 Antp intron Y Y(WEAK) Y Y 8 Hairy 5' intron Y Y Ubx 3' (and 9 NEW) Y Y(WEAK) Y 10 Ubx 5' Y Y 11 12 Ftz intron Y Y Y 13 Tailless intron Y Y Y 14 Bxd 3' Y(WEAK) 15 Bxd 5' Y Y 16 Sim P1 Y Y 17 Sim P2 Y Y 18 Sim 3' Y Y 19 CadN5' Y Y 20 CadN3' Y Y
20 LEGEND FOR NAMED PROBES
Gene DNP DIG Biotin Fitc Hairy Y Y Y Wg Y Y Y Caudal Y Y Y Giant Y Y Y Zen Y Y Y Cap n collar Y Y Y Tll Y Y Y Runt Y Y Paired Y Y Y Pnr Y Y AntpX (ncRNA) Y Y Y Y Scr Y Y Y Y Antp-P1 Y Y Antp-P2 Y Y Knirps Y Y Y Caudal Kruppel Y Y Y Hunchback Y Ubx 3’ Y Y twist Y Snail Y Y Y Ftz Y Y Y Hth Y Sim 3’ Y Y Eve 5’ Y Y
D. virilis Probes
Ftz Y Y Y Antp Y Y Scr Y Y Y
21
Some Gene descriptions:
Wingless (Wg) – segmental polarity gene Wingless is a lipidated secreted signaling molecule involved in early embryonic patterning as well as patterning of the wing, where it specifies the wing primordium and mediates the patterning activity of dorso-ventral compartment boundary
stage 4-5 stage 6-7 stage8-9
Cap’n’collar (cnc) – gap gene A Gap gene controlling some gene expression in the anterior of the fly. Activates segment polarity genes (like Wg and Hh) although the activation is not known to be direct or indirect.
stage 4-5
Caudal (c)– gap gene Expressed during early embryogenesis, distributed in a gradient with the highest levels in the posterior of the embryo. Shown to be essential for posterior hindgut formation and gastrulation. It is thought that caudal may target tailless and wingless amongst other things, as they are also required for some aspect of hindgut development.
stage 4-6 stage 7-8
22 Giant (gt)– gap gene Defines expression domains for pair-rule genes like even-skipped and paired. Hunchback represses Giant expression; caudal is also thought to control its expression. Giant also represses Hunchback and Kruppel to pattern the anterior of the embryo.
stage 4-6 stage 7-8
Hunchback – gap gene Central role in establishing an anterior-posterior gradient of gene activity in the early embryo. Post-fertilization zygotic Hunchback acts as a gap gene regulating Kruppel expression in the middle of the embryo and repressing knirps and giant expression so they are expressed only posteriorly.
stage 1-3 stage 8-9
Kruppel – gap gene Zinc finger transcription factor gap gene required later for proper generation of neural sublineages it is involved in Malpighian tubule bud morphogenesis; wing disc development; compound eye development
stage 4-6 stage 7-8 stage 9-10
Tailless – gap gene Expressed in the posterior of the embryo and involved in posterior patterning. Tailless is activated by the torso pathway.
stage 4-5 Hairy – pair-rule Hairy is a pair-rule gene expressed in a stripe pattern across the embryo. It is involved in forming segment identity.
stage 4-5 stage 6-7
23 Paired – pair-rule A pair-rule gene expressed in a stripe pattern across the embryo. Dissection of the paired promoter highlighted that all other pair-rule genes refined the expression of paired in the embryo.
stage 4-5 stage 6-7
Runt – pair-rule Runt, a primary pair-rule gene, is thought to modulate the expression of other pair-rule genes (for instance hairy, even-skipped). It also alters the expression of gap genes for instance Gt and Hb. It is expressed in 30 neuroblasts that give rise to the neuromere of the Drosophila embryo.
stage 4-5 stage 6-7 stage 8-9
Pannier – TF Also known as GATA-2, pannier is a TF. pnr is activated by decapentaplegic (dpp) in early development, but it has been found that after stage 10, the roles are reversed and pnr becomes an upstream regulator of dpp. This function of pnr is necessary for the activation of the Dpp pathway in the epidermal cells implicated in dorsal closure and is not mediated by the JNK pathway, which is also necessary for Dpp activity in these cells. In addition, pnr behaves as a selector-like gene in generating morphological diversity in the dorsoventral body axis. It is responsible for maintaining a subdivision of the dorsal half of the embryo into two distinct, dorsomedial and dorsolateral, regions, and also specifies the identity of the dorsomedial region.
stage 4-6 stage 7-8
Snail – TF Snail is a TF involved in Dorso/Ventral polarity. Twist and snail regulate invagination and commit cells to mesoderm fate. Snail alone is sufficient to intitiate the invagination of the ventral-most embryonic cells in the absence of twist gene activity. It acts as a transcriptional repressor, restricting neuroectoderm and neural fate in the invaginating mesoderm.
stage 4-6 stage 7-8 stage 9-10
Twist – DIG TF Twist acts to trigger mesodermal gene expression (for instance bagpipe).
stage 4-5 stage 6-7 stage 8-9
24 Zerknullt – DIG TF A transcription factor involved in DV-patterning. Although required only for a brief period (2-4hrs post- fertilisation), effects can be seen much later. For instance, in the absence of zen, germ band extension is abnormal.
stage 4-5
Antennapedia: hox gene
stage 4-5 stage 6-7 stage 8-9
Eve: pair-rule gene Even-skipped is a transcriptional repressor of a number of genes, including engrailed (acting indirectly through paired, runt and sloppy paired) (Fujioka, 1996), fushi tarazu, Ultrabithorax and wingless.
Bxd: long non-coding RNA
25 Single minded: transcription factor Single-minded is required for the developmental specification of the ventral midline. This is an organizing locus that appears as a result of gastrulation. Prior to gastrulation, there develop two anterior-to-posterior rows of single cells, one on either side of the embryo. All cells in both rows express sim.
stage 4-6 stage 7-8 stage 9-10
VII.3 Fixed Embryos :
Embryos: Wild type D. melanogaster (by age) D. virillis (limited numbers)
26 VII.4 Drosophila Stocks
Collection Type of Genotype Description type stock Cross Gal4 line Twi-Gal4 Gal4 expressed in twist domain Cross Gal4 line dpp-Gal4 (40Cb; FRT82B/CyO; Arol/TM6c) Gal4 expressed in Dpp domain Cross Gal4 line hh-Gal4/TM6b Gal4 expressed in Hh domain Cross Gal4 line nub-Gal4 Gal4 expressed in Nub domain Cross Gal4 line wg-Gal4 Gal4 expressed in wg domain Cross Gal4 line A9-Gal4 Expresses GAL4 in the wing and haltere discs Cross Gal4 line elav-Gal4 Gal4 expressed in neurons Cross Gal4 line Sp/CyO; dpp-Gal4 UAS-dcr2/TM6c Gal4 expressed in Dpp domain Collection Mutant Ubx[bx-3] Ubx[bxd-1] Ubx[pbx-1]/T(2;3)ap[Xa], ap[Xa] 4 winged flies (homeotic) Collection Mutant wg1, cn1 wg hypomorph (no wings) Collection Mutant cn[1] sli[2] bw[1] sp[1]/CyO slit (midline patterning defects) Collection Mutant wg(1-12) cn1 bw1/CyO wg amorph (segment polarity) Collection Mutant wg(1-17) Wnt2L/CyO wg amorph (segment polarity) Collection Mutant Ptc[9] cn[1] bw[1] sp[1]/CyO ptc (segment polarity) Collection Mutant Df (Kr)/cyo Kr (gap) Collection Mutant w[*]; lola[ORE119]/CyO lola (axon pathfinding defects) Collection Mutant y[1]; ftz[11] red[1] e[1]/TM3, Sb[1] ftz (pair-rule) Collection Mutant hb[12] st[1] e[1]/TM3, Sb[1] HB protein null (RNA is expressed) Collection Mutant w[*]; robo[2]/SM6b Robo (axon pathfinding defects) Collection Mutant Prd[9]cn[1]bw[1]sp[1]/cyo Prd (pair-rule) Collection Mutant w[*]; prd[9]/CyO; P{ry[+t7.2]=ftz/lacC}1 Prd (pair rule) Cross Other G-trace: UAS-dsred, UAS-FLP, Ubi>stop>nlsGFP G-Trace (II) Cross Other CoinFLP: eyFLP UAS-dcr2; CoinFLP UAS-GFP/CyO Mosaic Ga4/GFP expression in eye imaginal disc Collection Reporter nub-Gal4 UAS-GFP pouch of wing and haltere disks, rings on leg disks, salivary glands Collection Reporter btl-Gal4 UAS-act5:GFP Tracheal system Cross Reporter Moesin:GFP Moesin GFP (all cell outlines) Cross Reporter His2A-RFP Histone RFP (nuclei) Collection Reporter hh-Gal4 UAS-GFP/TM6b segmentation – wing disc (no expression before stage 7/8) Collection Reporter Vasa-GFP germline cells in embryo and ovaries/testes Collection Reporter UAS-dcr2; en-Gal4 UAS-GFP/CyO GFP expressed in En domain Cross Reporter UAS-Brainbow; elav-Gal4 Brainbow expressed in all neurons Cross Reporter UAS-Brainbow; rn-Gal4/S-T Brainbow expressed in rn domain (wing pouch) Cross Reporter UAS-Brainbow; hh-Gal4/S-T Brainbow expressed in Hh domain Cross UAS line UAS-mCD8-ChRFP Membrane Cherry Red Cross UAS line UAS-Kaede Photoconvertible GFP Cross UAS line UAS-Ubx/TM3 Ser UAS-Ubx expression Cross UAS line UAS-nls-GFP nuclear GFP Cross UAS line UAS-CIBN::Cre-N,UAS-CRY2::Cre-C light activable cre Cross UAS line UAS-ykiCA Constitutively active yorkie
27 Cross UAS line UAS-Cdk4, cycD Coexpressed Cdk4 and cycD Cross UAS line UAS-bantamC Bantam microRNA Cross UAS line UAS-InsRact Activated Insulin receptor Cross UAS line UAS-rasV12 Activated ras Cross UAS line UAS-Rheb Rheb Cross UAS line UAS-fatRNAi RNAi knockdown of Fat Cross UAS line UAS-spitfire Spitfire
Crosses
Genotype 1 Genotype 2 Description Moesin:GFP His2A-RFP Moesin GFP (all cell outlines) Histone RFP (nuclei) hh-gal4 GTRACE Lineage tracing nub-gal4 GTRACE Lineage tracing dpp-gal4 GTRACE Lineage tracing elav-gal4 GTRACE Lineage tracing twi-gal4 GTRACE Lineage tracing wg-gal4 GTRACE Lineage tracing A9-gal4 GTRACE Lineage tracing elav-gal4 UAS-mCD8-chRFP Nervous system labeled with membrane RFP dpp-gal4 UAS-Kaede dpp labeled photo-convertible GFP twi-gal4 UAS-nls-GFP cell nuclei of twist expressing cells nub-gal4 UAS-nls-GFP cell nuclei of nub expressing cells elav-gal4 UAS-ubx Mis-expression of Ubx in nervous system nub-gal4 UAS-ubx Mis-expression of Ubx in wing and haltere UAS-ykiCA CoinFLP Mosaic eye discs expresing UAS-X UAS-Cdk4, cycD CoinFLP Mosaic eye discs expresing UAS-X UAS-bantamC CoinFLP Mosaic eye discs expresing UAS-X UAS-InsRact CoinFLP Mosaic eye discs expresing UAS-X UAS-rasV12 CoinFLP Mosaic eye discs expresing UAS-X UAS-Rheb CoinFLP Mosaic eye discs expresing UAS-X UAS-fatRNAi CoinFLP Mosaic eye discs expresing UAS-X UAS-spitfire CoinFLP Mosaic eye discs expresing UAS-X UAS-Brainbow; elav-Gal4 UAS-CreN/C Multicolor labeling of neurons UAS-Brainbow; rn-Gal4/S-T UAS-CreN/C Multicolor labeling of wing imaginal disc pouch UAS-Brainbow; hh-Gal4/S-T UAS-CreN/C Multicolor labeling of hedgehog domain
28 VIII. DROSOPHILA DEVELOPMENTAL BIOLOGY & STAGING
1. Embryogenesis
Drosophila embryos complete embryogenesis in 22 to 24 hours at 25°C, and take approximately twice as long at 18°C. For the purposes of this lab, most of your fly experiments can be left on your bench, but embryo collections and GAL4 cross experiments should be done at 25ºC so that you can be sure of the developmental timing (refer to the Campos-Ortega and Hartenstein pictures for staging).
Gastrulation and segmentation are completed within the first several hours, and the last half of embryogenesis is mainly dedicated to organogenesis. Because development is so rapid, it can easily be observed under the compound microscope. The detailed study by Volker Hartenstein and José Campos- Ortega (see refs) remains the definitive description of embryogenesis, and this handout includes some of the staging tables and images from that study. Schematic diagrams from the fly atlas are also helpful in staging embryos.
Atlas of Drosophila Development Volker Hartenstein All embryos are in lateral view (anterior to the left). Endoderm, midgut; mesoderm; central nervous system; foregut, hindgut and pole cells in yellow. !!(amg) anterior midgut rudiment; (br) brain; (cf) cephalic furrow; (cl) clypeolabrum; (df) dorsal fold; (dr) dorsal ridge; (es) esophagus; (gb) germ band; (go) gonads; (hg) hindgut; (lb) labial bud; (md) mandibular bud; (mg) midgut; (mg) Malpighian tubules; (mx) maxillary bud; (pc) pole cells; (pmg) posterior midgut rudiment; (pnb) procephalic neuroblasts; (pro) procephalon; (ps) posterior spiracle; (po) proventriculus; (sg) salivary gland; (stp) stomodeal plate; (st) stomodeum; (tp) tracheal pits; (vf) ventral furrow; (vnb) ventral neuroblasts; (vnc) ventral nerve
29 Here is a schematic diagram that you can use to identify fly embryo developmental stages:
30 The pictures below (from The Embryonic Development of Drosophila melanoagaster by José Campos- Ortega and Volker Hartenstein) show what you will actually see if you cover the embryos with halocarbon oil and observe under the microscope, without removing the chorion. (You might also want to check out the Drosophila Atlas of Development by Volker Hartenstein, available at the “Interactive Fly” website: see URL p. 45)
31
32
2. Larval Development and Morphogenesis
After hatching from the egg, the first instar larva begins to feed and grow immediately. The first and second larval instars (L1, L2) last approximately 24 hours each, and the third larval instar (L3) lasts about three days. Groups of cells specified during embryonic development proliferate during these three larval stages, and form clumps of cells called imaginal discs in the lumen of the larva. These discs are the primordia of virtually all of the cuticular and ectodermally derived tissues in the adult.
Left: Fate map of imaginal disc primordia in a blastoderm stage embryo. Imaginal disc anlagen are represented as ovals. Right: Position of the imaginal disc primordia at the end of embryogenesis. Anterior is to the left. (ead) eye- antennal disc (wd) wing discs (ld) leg disc (hd) haltere disc (gd) gential disc
The imaginal discs of the eyes/antennae, head structures, legs, halteres, wings and genitalia are easily identified in L3, but are slightly more difficult to isolate from L2 and L1. Hox genes, among others, play important roles in specifying imaginal disc identity, such that specific cuticular structures develop on different body segments. The patterning of these discs during embryonic and larval development is the basis of pattern formation in the adult fly.
Left: Position of the imaginal discs in an L3 larva. Right: enlarged view of dissected imaginal discs and gonads of both sexes of an L3 larva. Anterior is up. Discs are shown in roughly the order that they appear in the larvae, anterior to posterior.
33
After the end of the L3 stage (approx. 5 days after egg laying (AEL)), the larva leaves the food and begins to crawl in search of a place to pupate. The anterior and posterior spiracles are everted, the puparium (pupal case) is formed, and the process of metamorphosis begins. Most of the larval tissue is destroyed by histolysis, but the imaginal discs continue to grow and undergo the morphogenetic movements that will form the adult fly. Dissecting pupae can be difficult since they are basically bags of mush with the imaginal discs floating around in them, but with practice it can be done.
Left: Colored coded scheme showing the imaginal discs with their respective adult structures. Right: Ventral view of a pharate adult within the pupal case prior to eclosion.
34 3. Adult Morphology
Most phenotypic markers used in standard fly genetics are dominant mutations that affect various cuticular structures in the adult. The position and morphology of different bristles, wing vein patterns, and other landmarks have been well characterized, so that you should be able to tell the difference between wild type animals and animals with specific phenotypic markers. Use the descriptions and drawings of mutant phenotypes from the red book (Genetic variations of Drosophila melanogaster), the Demerec book (Biology of Drosophila), and FlyBase (www.flybase.org) to help you learn to identify these markers.
35 4. The Female Reproductive System
Each of the two ovaries of an adult fly contains several ovarioles, which are compartments that look like a string of beads: the “beads” are oocytes at different stages of development.
Left: An adult ovary. Right top: A single ovariole. Right bottom: Picture of a pair of female ovaries still joined at the common oviduct, anterior is up.
The number of ovarioles varies slightly in different strains, but in our wild type strain the number is in the range of 15-20 ovarioles per ovary. Each ovary is covered by a thin peritoneal membrane or sheath, which is often rather impermeable and makes staining with many antibodies difficult, particularly the later stages of oogenesis.
The anterior of the ovary is called the germarium, which is a compartment containing germ line stem cells and somatic stem cells, encased in a covering of follicle (somatic) cells.
The germarium of D. melanogaster contains three cytologically disctint regions. Region 1 contains germ line and somatic stem cells; Region 2 contains mitotically dividing cystocytes; Region 3 contains the earliest stage of oogenesis.
These stem cells divide asymmetrically throughout the reproductive lifetime of the female, to produce oogonia and primary follicle cells. The oogonia (product of the divisions of germ line stem cells) divide
36 mitotically four times, and incomplete cytokinesis in each of these mitotic divisions results in a cluster of 16 cells connected by cytoplasmic bridges called Ring Canals.
Only two of the 16 cells are connected to the others by four cytoplasmic bridges, and one of these two always becomes the oocyte. It is thought that the quantity of a germ cell-specific organelle called the fusome that is inherited by both cells, determines which one of the two becomes the oocyte. The other 15 cells become nurse cells, and synthesise mRNAs and proteins that are delivered to the oocyte during oogenesis. Asymmetric localisation of many such mRNAs and proteins is responsible for the determination of the embryonic body axes and for the segregation of the germ line. The clone of 16 connected cells are of germ line origin, and are enclosed by a single layered follicular epithelium, which arises through mitotic division of follicle cells produced by the somatic stem cells in the germarium. These follicle cells undergo specific patterns of development and differentiation, include several different follicle cell types, and also play an important role in the patterning of the embryonic axes. As the oocytes mature, they move posterior in the ovariole, so that the most posterior egg chambers are the most mature.
Left: Schematic of a later stage oocyte. Right: A germarium and early oocyte stained with phalloidin and a ring canal-specific protein. Anterior is to the left.
37 5. The Male Reproductive System
The testes of adult males also have a stem cell region called the hub at the anterior, and the posterior regions of the testes are organized in only a rough chronological developmental series. Germ line stem cell divisions give rise to spermatogonia, which undergo four mitotic divisions followed by four meiotic divisions (one each). The resulting 64 spermatids are associated in a bundle, and remain clustered together during spermatogenesis.
From http://www.fly-ted.org/. Left: scheme showing spermatogenesis. Right: One half of the male testes false colored to show different regions and position of maturing sperm. Below: Dissected testes. Scale bar is 200µm.
38 6. Imaginal Discs and the Generation of Mosaic Tissues
Imaginal discs are epithelial sacks that proliferate extensively during the larval stages of development and differentiate during metamorphosis to form the adult structures such as the wing, eye etc. For the purposes of this course we will be focusing primarily on the wing imaginal disc (shown left). The wing disc precursors are set apart from the larval epidermis during embryogenesis. It is thought that each wing disc stems from an initial population of ~40 cells, which invaginate as an imaginal disc during early stage 17 of embryogenesis. These cells proliferate giving rise to a sack-like epithelial structure of ~50,000 cells, which consists of the squamous peripodial epithelium and the columnar epithelium of the wing disc proper. The basal side of both these epithelia face outwards to the larval body cavity, while the apical side is oriented inwards to the lumen.
Top: Drawing showing a side view slice through the developing wing disc. Compare to right-hand images below. (cu) cuticle, (ep) epidermis, (w) wing disc proper, (p) peripodial epithelium, (t) trachea. Middle: From McClure and Schubiger, 2005. Wing discs dissected from larvae at 12 hour intevals after egg deposition (AED). Images on left show a top-down (xy) view of the developing disc. Images on right show a z-slice through the same disc. Bottom: From Grusche, F. A. et al., 2009. Drawing showing normal orientation of wing disc, i.e. baso-lateral face of wing disc proper is “down” and apical face of wing disc proper is “up”. Blue dots = nuclei.
The wing disc gives rise to the adult wing, the wing hinge and a large portion of the adult cuticle, including the notum, scutellum and pleura. The wing disc proper forms the majority of the adult cuticle, while the peripodial cells form only ventral and lateral pleura (body wall) and ventral wing hinge.
39
Left: Fate map of a late larval wing disc. Yellow = notum and ventral pleura (body wall); blue = dorsal/ventral wing hinge; red = wing margin; green = wing pouch, light green will become the dorsal wing surface, dark green will become the ventral wing surface. Right: Dorsal view of corresponding adult structures. Ventral structures are not pictured. Wing discs, and others, are sub-divided into non-mixing, lineage restricted sets of cells called compartments. The earliest established compartment in the wing disc is the anterior-posterior (A/P) boundary, which is specified during embryogenesis and maintained throughout larval development and into the adult wing. Left: From Vincent, J.P., 1998. The wing imaginal disc anlagen is divided in two by a stripe of engrailed expression. Cells which express engrailed (yellow) will continue to do so and will assume an invariant posterior fate. Below: From Tabata, T., 2001. Wing disc and adult wing with posterior engrailed- expressing cells in blue. A second axis, the Dorsal/Ventral (D/V) boundary divides the wing disc pouch into the dorsal and ventral faces of the adult wing. The D/V border of the imaginal disc develops into the margin of the adult wing. A stripe of wingless expression runs along the D/V border.
40 The mechanisms that maintain these strict lineage restrictions are still not known. One hypothesis is that cells in different compartments have different affinities / adhesive properties and therefore segregate away from each other (like oil and water). Another hypothesis is that cells that enter an incorrect compartment are immediately signaled to undergo apoptosis – thus cell death might “sculpt” the smooth border. A limited number of mutations have been discovered that allow cells to cross compartment boundaries, but most of these pertain to the mechanisms that establish rather than maintain the boundary. For example, cells that do not receive the Hedgehog signal can cross from the anterior to posterior compartments because they do not attain anterior cell identity. The presence of the A/P boundary was established by use of a powerful technique in Drosophila genetics, mitotic recombination. This allows for the study of mutations that, when homozygous, might otherwise be lethal for the entire organism at an earlier stage of development. When a cell is heteroygous for a mutation, normal mitosis will result in the generation of two identical daughter cells that are each heterozygous for the mutation. In contrast, when mitotic recombination is induced, one daughter cell is homozygous for the mutation while its sister cell is wild type. If generated early in development, each cell generates a clone where all cells are identical. Thus, in a tissue such as an imaginal disc, where there is very little cell migration, mitotic recombination results in a clone of mutant cells adjacent to a wild-type sister clone that is often referred to as the “twin spot”. These days, mitotic recombination is induced using the FLP/FRT recombination system that has been adapted from yeast. FRT (FLP Recognition Target) is a sequence of nucleotides recognized by the FLP (Flip Recombinase) enzyme, which is native to yeast but also functional in D. melanogaster. The FRT sites can be introduced on chromosomes using transgenesis vectors such as P elements. FLP expression can be driven with ubiquitous heat shock or tissue specific drivers, in small or large groups of cells, throughout or at specific time points in development. This means that, with only a few exceptions, you can create groups of any number of cells, in any tissue, with mutations in any gene or combination of genes, at any time during development. The power of fly genetics in this respect remains unsurpassed by any other genetic model organism.
FLP/FRT- mediated mitotic recombination generates two genetically distinct daughter cells, either homozygous wild- type or homozygous mutant, from a heterozygous mother cell.
From Martin, F.A., 2009. Examples of clones within the wing imaginal disc. Note in the two left panels that the GFP- clones respect the A/P and the D/V boundary. In the right panel, clone and twin spot can be identified from the heterozygous tissue by either the lack of GFP or by carrying two copies of GFP.
41 The FLP/FRT system has been combined with the GAL/UAS binary gene expression system to produce powerful lineage tracing tools. GAL4 can be expressed under the control of a given promoter (e.g. constitutive promoters such as actin or tubulin or gene-specific promoters such as hedgehog or patched (see below)) GAL4 binds to Upstream Activating Sequences (UAS) and drives gene expression. To create lineage tracing tools, GAL4 is used to express FLP which then catalyzes mitotic recombination between FRT sites in cis. Unlike the trans FRT sites described above, which allows for sister chromatid exchange, FRT sites in cis allow for the excision of a small fragment of intervening DNA, which usually includes a STOP. This stop separates a constitutive promoter from your gene of interest, in this case eGFP. Therefore, in any cell where GAL4 drives FLP, GFP will be turned on and this expression will no longer rely upon continued expression of GAL4. The expression of GFP is stably inherited by all progeny of the original GAL4-epxressing cell. The figure below describes G-TRACE, a lineage tracing tool you will have access to in this course. Please see the original paper for more information: Evans, C. J. et al., Nature Methods 6, 603 - 605 (2009).
42
The Brainbow system in mice has revealed the morphology and connections of multiple interacting neurons in the brain, using Cre/Lox based recombinationLive imaging to Brainbow induce expression of different RESEARCH ARTICLE 1611 fluorescent proteins in different cells (Livet et al. 2007). This multi-cell labeling approach allows one to Fig. 8. Cellular dynamics during behavior. A crawling first instar larva image different cells as they exist and interact with exhibits rapid translocation and cell each other in vivo. Furthermore, this system can allow shape change of individual tendon 1606 RESEARCH ARTICLE Development 140 (7) cells (arrowheads), as visualized using one to distinguish and trace multiple clonal lineages in the GAL424B driver. The total time-lapse movie duration is 30 seconds. Scale parallel. bar: 20 µm. (Supplementary material Fig. 1. Molecular design. (A) Multicolor cell Movies 1 and 2.) Thelabeling. multi-cell ‘LOLLIbow’ labeling stock containsapproach a UAS- was recently adopted in Drosophilabrainbow transgene by several cassette groups encoding using color different genetic setups variants of GFP-like proteins: RFP (mCherry), YFP (Worley(EYFP) and et CFPal. 2013,(Cerulean) Boulina (top). Cre et al. 2013, Hadjieconomou et al. 2011,recombinase and Hampel stochastically1999; et Ritzenthaler al. removes2011). et oneTheal., 2000; of figures the Vasenkova below et al., are 2006). using In thethe developing cells have an assortment of means to exchange their own isolated wing epithelia, GPI-anchored GFP, a way to target GFP to molecules and that the multicolor stochastic cell labeling by “LOLLIbow”three lox cassettes system,the and, outer subsequently, which leaf of theyou plasmaonly will RFP, membrane,have access has furtherto in the revealed LOLLIbow would be useful in gaining new insights. courseYFP or (Boulina CFP can be‘argosomes’, et expressed al. 2013). which under The is the taken LOLLIbowcontrol as evidence for system intercellular is exchangebest suitedof GAL4 for (bottom). live imagingof ( stillB) Photo-inducible undetermined dynamic molecules ceCrell-cell (Greco interactions et al., 2001). The because pupal case it Simple genetics recombinase. ‘split-hasCre often’ stock been contains considered UAS- off-limits to intact imaging because of its To facilitate studies that incorporate our new imaging system usesCIBN::Cre-N bright,and spectrally UAS-CRY2::Cre-Copacity. Between separable, 24transgenes and 48membrane hours after puparium bound formation fluorescent (APF) further, we established viable homozygous stocks. First, we proteinsencoding (mCerulean, split fragmentsthe pupae EYFP, ofare Cre immobile,(top). and Blue mCherry).as they light histolyze larval tissues and form combined the two transgenes encoding the complementary halves the adult body. The combination of brightness and high expression of split-Cre into a single chromosome: UAS-CIBN::Cre-N,UAS- induces dimerizationlevels of of the the fluorescentcomplementary proteins facilitates direct image acquisition CRY2::Cre-C (the ‘split-Cre’ stock). This stock would require both Cellschimeric are markedproteinsof (bottom). assuperficial either ( Cepithelia, )mCerulean, Membrane such as the EYFP, wing, during or mCherry the pupal period. after GAL4 and blue light to initiate Cre activities. It allows for a control targeting reveals Furthermore,fine cellular detailsthe multicolor in each labeling cell of our LOLLIbow system over the cell type and its timing in which the DNA recombination a stochasticwith RFP, YFP recombination or CFPallows (cell visualization A, cell eventB, cell of C). involving two classes the of membrane-associatedCre occurs. Second, we combined the Brainbow transgene and the pan- recombinase and one of three pairs of variant loxP sites. Afterstructures this in event,intact epithelial this cellscell that, and by alldefinition, it’s progeny must arise from will neuronal GAL4 driver in one fly line: UAS-brainbow;elav-GAL4, continue to express the fluorescent protein. The timing of Cretheir neighborinduction cells ( ncan=27 moviesbe controlled in 20 pupae). byFirst, a the light- continually or the neuronal ‘LOLLIbow’ stock. Because the UAS-brainbow extending and retracting filopodia contact the surface of neighbors transgene has been integrated near either end of the second inducible dimerization of two halves of Cre (Cre-C and Cre-N).up to five The cells expressionaway (Fig. 9A). of Second, the fluorescentthe ‘argosomes’ ofproteins varied chromosome, it will be straightforward to create additional (and Cre) is under the control of a tissue specific promoterdiameters (promoter-Gal4). invade into the Thereforecytoplasm of neighborsone can (Fig. generate 9B). The ‘LOLLIbow’ stocks for various neuronal subpopulations or non- first occurs predominantly in the basal side, whereas the second neuronal cell populations by recombining other GAL4 drivers to multi-cell labeled tissue in any tissue and at any time in development.occurs frequently Additional in the apical side details of these cells.on the The mechanisms UAS-brainbow. Thus, the morphogenesis of individual neurons or LOLLIbow system can be found here: Boulina, M., Samarajeewa,for each, especially H., Baker, how they J. form D., andKim, what M. macromolecules D., & Chiba, they of other cell populations of choice in wild-type animals can now be may contain, need further assessment. However, it is clear that the examined with single-cell resolution in all the offspring from a A. (2013).Live imaging Live Brainbow imaging of multicolor-labeled cells in Drosophila. Development RESEARCH ARTICLEsingle1609 generation cross between ‘split-Cre’ and ‘LOLLIbow’ stocks and following a brief pulse of blue light illumination given at a desired time point. addition of dual palmitoylation sequence from GAP43 Movies previous work depended on cell labeling techniques that reveal (MLCCIRRTKPVEKNEEADQE). The construct was PCR amplified using Morphologically staged pupae were selected foreither imaging all synapses and studied indiscriminately using or only small subsets ofDISCUSSION them. PCR suppression strategy (Lukyanov et al., 1996; Luk’ianov et al., 1999) the same Zeiss confocal microscope or a LeicaWe SP5 propose laser-scanning that the confocal ‘image many, then analyze later’ approach,Five years ago, Livet et al. introduced their pioneering Brainbow and subcloned into the Omni vector. The construct was sequence verified microscope. Settings for the latter were adjustedwhich to minimize is possible exposure using time our system, could accommodatesystem swift to the neurobiology and developmental biology communities (Livet et al., 2007). Today, one thing still missing in prior to injecting into Drosophila. Transformants were generated using and laser intensity while still capturing high qualityscreens images: of cell-autonomous CFP and YFP and non-autonomous events during this and other topics of research in developmental biology. both fields, nevertheless, is vigorous application of this powerful attP16 and attP40 integration sites, respectively, on the second were detected with an Argon laser with 5 mW 458 nm (mCerulean) set to technology in experiments performed with live whole animals. The chromosome. 31% and 5 mW 514 nm (EYFP) set to 7%. RFP was detected with a 20 mW reasons for this omission might have been part technical and part Morphological details DPSS laser (mCherry) set to 9%. Dwell time was minimized by using the motivational in nature. We sought to push the methods for Cre recombinase Morphogenesis of individual neurons depends upon interactions resonance scanner at 8000 Hz. Data consist of z series through the 10-15 µm visualizing the dynamic cellular processes directly, with single-cell To generate Split-Cre constructs, we essentially used the same vector and with neighboring neurons. One area that has rarely benefited from epithelium, collected over time. The movie of a crawling first instar larva, resolution, within intact organisms throughout their different stages strategy. Nuclear targeted CRY::CreN and CIBN::CreC chimerical ORFs direct visualization in vivo is the interactions among neuronsof ofdevelopment. the on the other hand, used a spinning disc confocal microscope with a 10×/0.30 (Addgene ID 26888 and 26889) were separately cloned into Omni vector same molecular identities. Genotype-independent, stochasticAll images in this paper were from live unfixed specimens of Plan Neofluar lens. This system consisted of a Yokogawa CSU X1 spinning using XhoI and XbaI cloning sites and the following primers: multicolor labeling by Brainbow allows for many cells of thewild-type same Drosophila. They were imaged using confocal aaaCTCGAGatgaatggagctataggagg (CIBN_XhoI_DIR),Fig. 3. Tricolor aaaTCTAGA labeling.- Larvaldisk brain unit, labeled equipped using elav-GAL4 with continuousdriver: wavetype lasers, to be dichroic visualized filter individually sets with distinct colors (Livetmicroscopy et al., at various developmental stages. A laser-scanning fluorescent microscope equipped with photomultiplier tubes was ctaatcgccatcttccagca (Cre_C_XbaI_REV), aaaCTCGAGatgaagatggac(A) default expression -(RFP) (Semrockwithout recombinase Di01-T 405/488/568/647 activities; (B) tricolor beam splitter2007). with The 483/32 fine and details 542/27 of individual neurons labeled with the labeling (RFP, YFP or CFP) withemission recombinase filters activities. for 488 Bothnm andOFP and561 RFPnm lasers, respectively) and an EM- used to collected most images, while one movie that captured fast aaaaagac (CRY2_ XhoI_DIR) and aaaTCTAGAttacagcccggaccgacgat stochastic Brainbow system can be visualized best by using cellularepitope behavior was acquired using a CCD equipped spinning disc are expressed as default (seeCCD text). cameraScale bar: (Photometrics 10 µm. Quant EM 512SC) mounted on an inverted (Cre_N_XbaI_REV). Vectors were integrated using phiC31 into attP16 and tags that permit nearly unlimited signal amplification (Livetconfocal et al., microscope. However, any conventional confocal system attP40 sites on the second chromosome. The construct was sequence Zeiss AxioObserver fluorescent microscope2007; (Intelligent Hampel et Imaging al., 2011). and Unfortunately, this method is notcapable readily of spectrally separating the signals from the fluorescent Innovation). Exposure time for each channel was 500 mseconds. All movies verified prior to injecting into Drosophila. make the situation better. Increasing the copy number of the Fig.compatible 9. Cellular dynamics with during live imaging.wing formation. An ( alternative,A) Filopodia (arrows) live imaging-proteins used in our system should be able to provide similar results Brainbow cassette from onewere to imaged two, however, at room produced temperature. six distinct extendcompatible and retract actively method to reach would neighboring be to cells. create, (B) Argosomes within each (see cell, Materials a and methods). To preserve fluorescence of cell labels Induction of Cre colors instead of three (Fig. 4). Knowing the exact copy number of (circles)concentration diffuse around in gradient the cytoplasm of of fluorescence neighboring cells. towards Scale bar: the as plasma well as the health of the living samples, a minimal number of Live imaging detection channels and illumination time were employed in our We used a mercury lamp (X-cite 120PC Q 120 W mercurythe cassette vapor short within arc the genome permitted an unambiguous color 10 µm.membrane. (Supplementary Both material manually Movies applied 1 and 2.) carbocyanine fluorescent43 dye Before imaging the live embryos, we removed the chorion membrane lamp) and a GFP filter (FS 38HE filter configurations:separation. excitation This 470/40, could be useful, especially when tracing individual DiI (Honig and Hume, 1986) and genetically delivered chemically with 50% bleach solution exposure for three minutes at room dichroic 495, emission 525/50) to induce recombinase activitiesneurons with of the long split- axons that span across other neurons and tissues. palmitoylated GFP (Moriyoshi et al., 1996) exemplify previous temperature. Embryos were transferred onto a large 40ϫ22 mm coverslip Cre. Embryos expressing the complementary Cre halvesMulticolor received a labeling full field of cells could also influence the experimental approaches that attain single-cell resolution without fixation of illumination through a 10×/0.30 Air Plan Neofluar design.objective One lens. goal The of experiments(No 1.5 from using Corning GAL4 driversor VWR), is to covered isolate withtissues. halocarbon The effectoil (Halocarbon of membrane-targeted fluorescence is, in fact, illumination pulse of a varied duration consisted of alternatingevents that 20 take seconds place withinProducts, cells CAS#9002-83-9),of a discrete population. and mounted Here, on themost inverted striking microscope. when one visualizes For neurons and non-neuronal cells ON followed by 20 seconds OFF (Fig. 2A). It should bethe noted typical that logicsamples is as follows:imaging the the live smaller larvae, the their population cuticle skin size was washedknown thoroughly to extend verywith smallwater. and extremely fragile structures, such were exposed to normal room light. examined, the more straightforwardExcept for becomes taking moviesthe interpretation to capture of the its locomotionas filopodia, of theduring sample development (see (Ritzenthaler et al., 2000; Kim et data. With multicolor labeling,supplementary however, material the strategy Movies changes, 1, 2), we i.e. used anal., extra 2002). coverslip Because (18×18 chemical mm and physical treatment are not required Microscopy use color separation andor simultaneously 22×22 mm number watch 1 or events 1.5) over in athe large vacuumprior grease to spacerimaging, and data with collection some is accelerated, whereas issues of Confocal imaging number of cells, then seekmanually impacts ofapplied interactions pressure among to minimize cells of thetheir movementfilopodia during retraction the imaging and fluorescence loss are minimized. Confocal images were taken on a Zeiss 780 inverted confocalsame and/or microscope different populations.session. The For sufficient example, pressure combining was individually both Interactions checked carefully among in neurons each that posses equal competitive with Zen 2010 software, using a 40×/1.3 Oil Plan Neofluarelav-GAL4 lens. Immersoland GAL24B driverscase though permitted visual simultaneous inspection, visualization while the totalcapabilities imaging duration have been was an kept area of intense research. Here, because of 518F from Zeiss was used for immersion with the latter.of three A 25 motoneuron mW Argon endingsminimal that areat 5-20co-innervating seconds per a single512×512 muscle pixel slice.its compact Pupae were size, each the washedspace in which intercellular interactions take laser was used as the source for the 458 nm (mCerulean),along 488 with nm nearby (EYFP) tendonand cells mounted within ina asingle glass-bottomed animal (n=6 25 larvae,mm Petri dishplace (WillCo-Dish). becomes small A layer in Drosophila of , enhancing the chance that and 514 nm (Kusabira Orange) excitation and a 20 mWFig. DPSS 5). Inlaser the for larval the neuromuscularthiodiethylene system, glycol the between motoneurons the pupal of caseindividual and the coverslipneurons of reduced equal molecular capacities can potentially 561 nm (mCherry) excitation. Laser powers of 9.0%, 3.5%,molecularly 1.2% and and 1.2% functionallyrefraction. disparate Following properties the controlimaging the session, entire samplestouch were one each another. collected For example, with bilateral neurons present on both with spectral detection ranges of 463-500 nm, 498-515set ofnm, skeletal 550-568 muscles nm attacheda soft brush to corresponding and transferred tendon onto yeast-agar cells and plates.sides of the brain extend their axonal and/or dendritic growth cones and 625-669 nm, respectively, were used as acquisitionsurrounded settings with by a pixel other cells of different functions. In order to reciprocally. They would, therefore, encounter an identical dwell time of 50.4 µseconds. Where appropriate, tilingcharacterize and digital zooming both geneticDrosophila and activity-dependentstocks mechanisms that molecular environment twice, i.e. before and after crossing the were applied. No spectral unmixing was employedcontribute to distinguish to the the developmentThe following of each homozygous neuromuscular stocks synapse, are availablemidline upon(Furrer request: et al., 2003).UAS- Peripheral sensory neurons that occur multicolor labels. CIBN::Cre-N,UAS-CRY2::Cre-C (UAS-CIBN::Cre-Nin reiteratedat patterns attP16 could and also see their molecular environment repeated across the segmental border. Some such neurons are known to exhibit self-avoidance when establishing complex dendritic arborization patterns (Grueber et al., 2002; Grueber et al., 2003). GAL4 drivers are, by nature, incapable of distinguishing individual cells among molecularly identical cells. As a result, simultaneous visualization of these and other cases of intercellular interactions among bilateral and segmental homolog neurons have previously relied on labor-intensive dye injection and/or computer-assisted tracing of individual neurons. LOLLIbow, however, was able to unveil the fine dendritic morphology of individual segmental homolog neurons in a live larva with distinct colors (n=4 third instar larvae, Fig. 6). Studies characterizing how various molecular signals sculpt neurons could benefit from the single-cell resolution and multicolor imaging of neurons that are molecularly identical and yet interact extensively with one another within an animal.
Fig. 4. Color variations. Tendon cells colored using the GAL424B driver: Live imaging (A) three colors with single copy of the Brainbow cassette; (B) six colors Understanding the mechanisms that regulate the morphological with double copies. Scale bar: 10 µm. dynamics of cells is fundamental to developmental biology. In the The combined FLP/FRT and Gal4/UAS system remains an efficient and effective method to generate mosaic tissue in drosophila. One can easily generate patches of cells expressing Gal4 in a background of wildtype cells, which then activates transcription of your gene of interest in these patches only. This tends to generate patches of overexpressing cells throughout the animal, which for some potent genes (such as oncogenes like activated Ras) can cause animal lethality or delay development. Furthermore, generation of the mosaic tissue requires a heat shock (to induce FLP expression), and the developmental timing and length of FLP induction can greatly influence how many cells excise the FRT stop FRT cassette and activate Gal4.
The Hariharan Lab has built a genetic system called “CoinFLP”, which is a modification of the combined FLP/FRT and Gal4/UAS system. The CoinFLP system automatically generates mosaic tissue expressing Gal4 in a specific organ (for example, the eye imaginal disc when using eyeless-FLP) and produces similar proportions of Gal4 expressing cells from animal to animal. The main advantage of the CoinFLP system is the ability to quickly analyze mosaic tissue expressing many different UAS- constructs (e.g. gene overexpression or knockdown), facilitating large scale unbiased screens with hundreds of UAS lines.
Similar to the Brainbow system, the CoinFLP system uses FLP recombination of two pairs of variant FRT sites. Recombination between canonical FRT sites results in the excision of a STOP cassette such that the Gal4 gene can be transcribed and translated. Thus GAL4 is expressed in that cell and all of its descendants. In contrast, recombination between the two FRT3 sites results in the excision of the one of the canonical FRT sites thus preventing any subsequent FLP/FRT-mediated recombination while maintaining the STOP cassette upstream of GAL4. Thus this cell and all its descendants cannot express GAL4 protein. The FLP recombinase is expressed under the control of the eyeless promoter thus restricting recombination events to the eye-antennal disc. In order to examine the phenotype of imaginal discs, CoinFLP can be used in conjunction with UAS-GFP. We will use this CoinFLP UAS-GFP system in class to examine the effects of mosaic expression of genes that positively regulate growth, which can cause overgrowth of the GFP tissue.
44 Arthropod Lab Part 2: Other Arthropods
Parhyale
Live Imaging of Parhyale Embryos Several aspects of Parhyale development lend themselves to live-imaging experiments. For example, the lineages of the early blastomeres are restricted, making them well suited for lineage tracing and ablation experiments. Additionally, we have brought two transgenic lines that will facilitate your observations of Parhyale development. We recommend you begin with injecting one-cell embryos before moving up to two- through eight-cell embryo injections. You may visualize Parhyale development by filming your tracer-injected embryos, or one of the Ds-Red transgenic lines.
Reagents available for Parhyale live imaging: Labeling Reagent/ Usage Information Antibody/ Notes Transgenic Line (Live Embryos) Fixed Embryos FITC-Dextran 50mg/ml stock Anti-Flourescein Green fluorescent dye will allow (Fluorescein isothiocyanate- Use at 10-205 mg/ml for Use at final conc. of you to visualize your embryo dextran) tracing, and >25mg/ml 1:3000 immediately. (Note, adding too for ablations much will kill the cell almost immediately.) To ablate cell lineage later in development, shine blue light on FITC-injected cells. HSP-NLS-DsRed Parhyale Heat shock embryos: n/a Nuclear-localized red protein- after (transgenic line produced by 1 – 1 ½ hour at 37°C/day heat shock, will appear in all cells. M. Modrell, HS promoter from T. Pavlapoulos) Muscle-DsRed Parhyale A muscle promoter drives n/a Red protein in muscles (transgenic line produced by DsRed in this line. R. Parchem, muscle promoter Sit back and watch! from T. Pavlapoulos)
Parhyale RNAi injections
You may probe the function of the single-minded and abdominal-B genes by knocking down their function with sim or Abd-B siRNA injections.
The sim phenotype is apparent 4 – 5 days post-injection. Look for the two pairs of long antennae before dissecting. Sim affects the ventral midline, and severe phenotypes are radialized and lack all appendages but the first and second pairs of antennae. Fixing and dissecting these embryos is fast and simple using the boiling fixation method. The boiling method should only be used prior to the leggy stages, because the embryo won’t unfold and lay flat after boiling. (Ask Heather for boiling method protocol if this is of interest).
The Abd-B phenotype transforms the pleopods (swimmerets, A1-A3) to periopods (walking legs). It’s a pretty dramatic phenotype! Note that the phenotype isn’t apparent until very late in embryogenesis (best observed in hatchlings, about 11 days), so you will have to complete your observations after the arthropod section is over.
Parhyale embryo Fixation and Dissection
Being patient is the most important part of embryo dissections. A small number of embryos that have been dissected and fixed well may be more valuable than numerous embryos in pieces.
Extracting embryos from females using clove oil: Gravid Parhyale females brood their embryos in a ventral pouch (See Section VIII.1, white arrow in lower panel (b) of first figure). To extract embryos without sacrificing the mother, you can put your female Parhyale to sleep using clove bud oil in seawater. **It is especially important to put your females to sleep if they are transgenic, you want to be able to use these females again!** 1. Putting amphipods to sleep: a. Add 10uL of clove bud oil to 50mL of filtered seawater in a falcon tube. Shake vigorously. b. Collect gravid Parhyale in a Petri dish or a medicine cup. c. Remove as much water as possible. d. Add your clove oil / seawater mixture (cover the Parhyale). e. Wait for them to completely stop moving – 5 to 10 minutes should do the trick. f. Caution: Leaving amphipod in the clove oil too long (hours) will kill them. 2. Embryo extraction: a. After the Parhyale are asleep, transfer them to a sylgard plate using forceps or a plastic transfer pipette (with tip cut off so amphipod will fit through opening). b. With the forceps at a steep angle, corral the waist of the animal, i.e. between the forward- and rearward-facing walking legs. Hook the glass probe between the walking legs and rotate the animal ventral side up. Grasp the waist of the animal until the legs splay just a bit. They’re slippery, so it will take some practice. c. Use the small blunt end of the probe [to make probe see additional protocol] or forceps to sweep out the embryos by starting at the posterior end and moving the probe through the brood pouch. d. ** Be careful not to damage the embryos - the younger animals are very soft and are squished easily! Also, do not damage the females, especially the transgenic ones!** e. Transfer embryos to a new petri dish or tube. Wash with seawater to remove clove oil. f. To wake mother amphipods up, remove them from the clove oil and place them in a dish or cup of clean seawater until they recover. Return clove oil mixture to a Falcon tube – this can be reused. g. Put the adult amphipods back into their tank. Note: Do not place sleeping amphipods back into their tank – they will be eaten by the others in the tank!
Any embryos removed from their mothers should be stored in filtered seawater and placed in a humidity chamber (a.k.a. pipette tip box lined with wet paper towels) on a bench top, or a 26 degree incubator.
1 Dissecting and fixing Parhyale embryos You will need: Forceps (optional) 3 well glass dish or Plastic Petri dishes (one with Sylgard) Dissecting needles Medicine cups (optional) 3.7% formadehyde in filtered seawater PT to rinse fixed embryos Eppendorf tubes Glass Pasteur pipettes and/or plastic transfer pipettes Helpful setup tips: • In the 3-well glass dish, fill the first well with filtered seawater, this will be your “corral” of embryos waiting to be fixed; fill the middle well halfway with fix, this is where you’ll fix and dissect simultaneously; and fill the last well with filtered seawater, this will be where you rinse the fix off of your dissected embryos. • Embryos tend to stick to glass pipettes and even the sides of the wells in the glass dishes. An easy way to prevent sticking is to use some yolk/material from the first group of embryos you dissect to swirl around the bottom of the dish, also use to coat the insides of your transfer pipette prior to using with fixed intact embryos. Only pull embryos up into the narrow neck of the pipette – avoid the expanded upper region of the transfer pipettes, embryos tend to get trapped there.
Protocol: 1. Place a few embryos (start with 2-3 and increase with experience) in a dish containing fixative. 2. Holding each embryo in place with your forceps or one of your dissecting needles (forceps are optional), poke a shallow hole in the eggshell with a dissecting needle. A shallow hole will avoid damaging embryonic tissues, and try to poke a hole in the yolk away from most of the embryo if possible (see figure next page). This is tricky for very early embryos because their cells are evenly distributed around the yolk (the cells of older embryos condense to one side). Start your timer (or note the time on the clock) after you have poked a hole or made some kind of tear in each embryo in fix. 3. Allow fix to enter the embryo for a couple of minutes – this assists in the dissection – however do not wait too long because the embryo is also fixing to the outer membrane(s). 4. Carefully peel away the outer egg shell membrane starting at the hole you made with your dissecting needle. Sometimes it helps to make a slight tear in the shell at the poked hole because it will produce a flap or loose end that you can hold with a needle or forceps. 5. While holding a piece of free membrane with one needle, carefully peel the membrane away from the embryo with the other needle. Sometimes it may be easier to hold a piece of membrane and try to roll the embryo away from the membrane with another needle. Either way, it is important to remove the membrane from the tissue as gently as possible. Be careful of appendages sticking to the membrane – it is very easy to dismember the embryos. 6. If you are lucky, the inner membrane (germband stages and older) will come off with the egg shell membrane! If not, repeat steps 4 & 5 for inner membrane. 7. Try to get the embryo mostly out, and exposed to fix relatively quickly – the time it takes you to remove the membrane will influence the amount of time you actually fix the embryo. Try to remove the membranes within 5-10 minutes and then allow the embryos to continue to fix for 10 more minutes (total time in fix should be around 15-20 minutes). 8. Once fixed, carefully pipette embryos into a dish/tube containing PT to rinse embryos before beginning the staining protocol.
2 General things to keep in mind while dissecting: Be careful. Don't make any sudden or jerking movements, this will tear the embryo. Be patient – large pieces of embryos will also give you some data. You do not have to remove every bit of yolk from the embryo when antibody staining as long as enough yolk has been removed to expose the tissue sufficiently, your staining will work fine.
Helpful hints for dissecting different age Parhyale embryos: 0-18hrs: These embryos are very yolky and it is difficult to maintain overall shape while dissecting away the single membrane surrounding them. Poke a very shallow hole to begin. You may want to initially fix for a few minutes while you dissect and start the real time of fixing once you have totally removed the membrane. For example, you may want to spend 5 minutes in seawater plus formaldehyde (9:1) to poke a hole and remove the membrane followed by another 15 minutes in fresh/another fixative (9PEM:1PBS(10x):1Formaldehyde) once the membrane has been removed.
1-2 day: These embryos are relatively easy to dissect because they are surrounded by a single membrane and since most cells have condensed to one side of the embryo, you can pretty much hack off the yolky side of the embryo without losing too many cells.
60hrs - 3days: This is another tricky stage because the embryo is not easy to identify and the yolk has developed two membranes – one of which is easy to remove (outermost) and another inner membrane that is more difficult to remove. You still want to poke a hole in the embryo to begin your dissection. One good way to locate the embryonic tissue is to roll the embryo on its side in the well with fix. As it rolls around, you will notice an arc of whitish or more opaque region with respect to the purplish yolk and totally clear space sometimes seen between the membrane and the yolk. Pierce the embryo on the opposite side from the opaque region. The only other hint at this stage is to dissect fewer embryos at a time – therefore you can remove both membranes quickly before the inner one sticks to the embryo like plastic wrap. The longer the embryo sits in fix, the more the inner membrane becomes fixed to the embryo. If you cannot remove all the membrane, remove all the yolk on the opposite side – this exposure is generally enough for antibody staining to work, however a through removal of the inner membrane is best for good in situ results.
3 4+ days: Once the embryo has grown appendages, the trick is to remove the membrane without breaking off appendages. It is easier to dissect older, leggy embryos if you poke a hole dorsally just posterior of the head. This will give you more room to begin removing membranes. After poking a hole it can be helpful to let them sit for 1-2 minutes undisturbed in fix.
Phalloidin Staining (muscle stain) Note: Embryos must not have been exposed to methanol! 1. Fix in 3.7% formaldehyde 20-30minutes at RT 2. Wash 2X with PT 5-10 minutes* 3. Wash 10 minutes with 70% cold acetone 4. Wash 10 minutes with100% cold acetone 5. Wash 2X with PT 5-10 minutes (if your hatchlings are more than a few days old, sonicate a few seconds) 6. Incubate 1:500 overnight at 4 °C (a couple of hours at RT will also work) 7. Rinse with PT 3X 8. Stain with DAPI/50% glycerol, if desired 9. Store in 70%glycerol *If you want to stain with phalloidin and an antibody, perform the antibody stain first.
4 Parhyale development and staging (Modified from Browne et al (2005) Genesis 42:124-49 and Gerberding et al, (2002) Development 129:5789- 5801.) A. Introduction Studying the relationship between development and evolution and its role in the generation of biological diversity has been reinvigorated by new techniques in genetics and molecular biology. However, exploiting these techniques to examine the evolution of development requires that a great deal of detail be known regarding the embryonic development of multiple species studied in a phylogenetic context. Crustaceans are an enormously successful group of arthropods and extant species demonstrate a wide diversity of morphologies and life histories. One of the most speciose orders within the Crustacea is the Amphipoda. The embryonic development of a new crustacean model system, the amphipod Parhyale hawaiensis, is described in a series of discrete stages easily identified by examination of living animals and the use of commonly available molecular markers on fixed specimens. Embryogenesis is completed in approximately 250hrs at 26°C and has been divided into 30 stages. This staging data will facilitate comparative analyses of embryonic development among crustaceans in particular, as well as between different arthropod groups. In addition several aspects of Parhyale embryonic development make this species particularly suitable for a broad range of experimental manipulations.
5 B. Reference Guide to Parhyale Development 1. S1-4 Oocyte to eight cell stage, lineage of eight cell stage, 0-9hrs of development Early cleavages are total or holoblastic, resulting at the eight cell stage in an embryo with 4 micromeres and 4 macromeres. The lineages of these early blastomeres are restricted early in development such that the mesoderm is derived from only 3 blastomeres: ml (mesoderm left side), mr (mesoderm right side) and Mav (anterior and visceral mesoderm), while the ectoderm is derived from the El (left), Er (right) and Ep (posterior and midline) blastomeres.
2. S6 Soccerball stage Cells are approximately the same size at this point; the divisions are asynchronous and the yolk is shunted internally to center of embryo.
3. S7-S8 Rosette stage - Gastrulation The rosette, which is made up of Mav and g progeny, marks the future anterior side. The ectoderm will migrate ventrally, and then over the rosette and mesoderm progeny. The rosette is no longer visible by S8. After this the germdisc continues to condense on the anterior ventral side.
6 4. S9-17 Germband formation and elongation Ectodermal and mesoblast rows are organizing along the ventral surface in transverse rows. The midgut anlagen is visible as an aggregate of cells on either side of the head lobes that becomes more organized as an ovoid anlagen (triangles). By S17, the caudal furrow is visible at the posterior (arrowhead) and the germcell cluster has split into bilateral clusters (arrowheads). Limb buds are developing on the anterior region of the animal. On the right is a series of dissected embryos stained with the segment polarity gene, Engrailed and counterstained with a nuclear dye, DAPI, showing the progression of segmentation along the A/P axis.
Left panel: live image and matching DAPI (nuclear) images, Right panel: ventral view of embryos stained with Engrailed Brightfield and DAPI images shown
7 5. S18-30 Appendage formation, organogenesis and neurogenesis to hatching At these stages, the posterior regions (telson), gut and limbs become well developed. The germcells migrate to a lateral position between the ectoderm and midgut at S21 (arrowhead) and then by S28 have migrated dorsal medially as the embryo undergoes dorsal closure. The hindgut proctodeum (arrow at S21) is visible at the posterior terminus and digestive cecum begins to extend posteriorly (arrow at S24). Eye fields and a beating heart also begin to form by S28, followed by cuticle thickening and muscular twitching before hatching at S30 (250hrs)
Left panel: live image and matching DAPI (nuclear) images, Right panel: ventral view of animals stained with Engrailed, Brightfield and DAPI images shown.
8
V
9 Artemia
These crustaceans require special permeabilization techniques for optimal staining. You only need a few embryos per staining (but you can have the equivalent of 20 µl packed volume if you want). It is a good idea to throw a couple of fly embryos (50 or so; rehydrated in PT) into the same tube to act as an internal control.
Fixation: 1. Fix animals in 9:1:1 FSW OR PEM:10X PBS:32% formaldehyde (3.7% formaldehyde in 1X PBS works fine as well) for 17-20 minutes at room temperature. Agitate the tube gently during fixation. 2. Rinse animals in PT several times (5 or 6 changes). 3. Proceed directly to permeabilization and staining or dehydrate into methanol gradually and store in 100% methanol at –20C.
Rehydration: 1. Rehydrate animals gradually into PT. Remove 1/4 of the methanol in the tube with the animals and replace with PT. Repeat this 5-6 times. 2. Rinse the animals in 100% PT several times (5 or 6 changes).
Permeablize and Block: 1. Wash animals 2 X 10 minutes in PT. 2. Permeabilize cuticles by sonication or detergent treatment. We will use sonication only. The detergent recipe is provided for your information. a. Sonication: Give 2 to 4 brief pulses (about 3 seconds) in a bath sonicator (see T.A. for help). Dip them in three or four times and pull them out each time when the tissue swirls around and clumps together to form a little pulsating ball. b. Detergent treatment: Incubate animals in a solution of 0.3% Triton-X + 0.3%sodium deoxycholate at room temperature for 30 minutes to 1 hour. 30 minutes is sufficient for young animals. Increase time in detergent for older, larger animals. 3. Wash animals with PT 5 or 6 times then leave in PT for 15 minutes. 4. Block animals in PT+N (PT + 5% NGS) for 30 minutes.
Antibody Incubation: Follow the previous general protocol for Drosophila with the exception that you may want to incubate tissue in appropriate dilution of secondary antibody (dilute antibody in PT+N) overnight at 4°C.
Clearing: 1. Wash overnight in PT to reduce background. 2. Clear tissue in 50% glycerol + 1µg/ml DAPI for a few hours at room temperature or overnight at 4°C. 3. Clear tissue in 70-80% glycerol overnight at 4°C until the tissue sinks. Tissue is now ready for mounting and photography.
Mysids& ! Mysids!develop!within!the!brood!pouch!of!the!female!in!four!days,!corresponding!to! Stages!1;4.!!Whole!females!with!embryos!are!fixed!with!3.7%!PFA,!stored!in! methanol,!and!when!needed,!can!be!rehydrated!in!PT.!!Each!female!carries!three!to! six!embryos.! ! Stage&1:!Yellow!ball.!!Corresponds!to!the!germband!stage.!!Square!cells!are! organized!into!perfect!rows!and!columns.!!The!eggs!are!easily!removed!by!grasping! the!female!with!forceps!and!brushing!the!brood!pouch!with!another!pair!of!forceps.!!! The!embryos!must!then!be!must!be!dissected!out!of!the!vitelline!membrane!with! forceps!or!tungsten!needles.!!Do!not!sonicate!this!stage.!!Embryos!hatch!out!of!the! vitelline!membrane!following!Stage!1,!so!all!subsequent!stages!do!not!need!to!be! dissected.!!! !
!! !! ! (left)!A!female!with!Stage!1!embryos!next!to!four!Stage!1!embryos.!(middle)!Cluster! of!stage!1!embryos.!!(right)!Stage!1!embryo!with!membrane!removed.! ! ! Stage&2:!Tadpole.!The!embryo!has!legs,!but!they!are!held!tight!against!the!body,!so! they!are!difficult!to!see.!Stage!2!embryos!stick!together!and!must!be!gently!pulled! apart.!This!stage!may!not!need!sonication,!but!can!be!gently!sonicated!to!open!up! the!cuticle!for!weaker!stains.! !
!! !! ! !(left)!A!female!with!Stage!2!embryos!next!to!four!Stage!2!embryos.!(middle)!Cluster! of!stage!2!embryos.!!(right)!Separated!stage!2!embryos.! ! Stage&3:!White!eyes.!The!embryo’s!legs!are!now!visible,!and!it!has!white!eyes.!This! stage!must!be!sonicated.! !
!! !! ! (left)!Female!with!Stage!3!embryos!next!to!three!Stage!3!embryos.!(middle).!!Brood! pouch!with!Stage!3!embryos.!!(right)!!Isolated!Stage!3!embryos.! ! Stage&4:!Red!eyes.!The!embryo!has!red!eyes!and!more!detailed!features.!This!stage! must!be!sonicated.! !
!! !! !!! (left)!Female!with!Stage!4!embryos!next!to!three!Stage!4!embryos.!(middle).!!Brood! pouch!with!Stage!4!embryos.!!(right)!!Isolated!Stage!4!embryos.! ! ! Sonication:!!Place!embryos!to!be!sonicated!in!1.5!ml!eppendorf!tubes!in!1!ml!PT.!! Stage!2:!sonicate!40;60!seconds.!!Stage!3!and!4:!Sonicate!15!minutes.!!All!sonication! in!Branson!1510!water!bath!sonicator!located!in!cold!room;!put!eppendorfs!into! floaties!to!sonicate.!!After!sonication,!rinse!with!PT!and!begin!antibody!staining!as! you!would!Drosophila!(block,!add!primary,!etc.).! ! Spiders (Remove the eggs from the egg sac before you start this protocol.)
Dechorionate spider embryos in 50% bleach in PBS, 5-10 minutes. The best way to do this is to add 50% bleach to the dish with the spider embryos and then “dive bomb” them with a Pasteur /transfer pipette until they remain submerged. Agitate occasionally to remove bubbles from the surface of the embryos. **After they are submerged, keep them under water throughout the rest of the protocol, otherwise, they stick to each other** 1. Rinse. Transfer embryos to an eppendorf tube and rinse carefully with 1x PBS. Five quick washes should remove all the bleach. 2. Fix and Dissect. Use 9:1 1XPBS: 32% formaldehyde. It is a good idea to poke holes in the embryo and allow it to sit for 5-10 minutes before pulling off the membrane and fixing for another 10 minutes (15-20min total in fix). Rinse and stain or dehydrate and store in MetOH.
Stages in the embryogenesis of Zygiella x-notata. Development takes roughly 250 hours at 22ºC. Embryos oriented posterior down in all panels. Time given in hours after egg laying A–H, View of germ disc side. I–O, Ventral view. A, Pre-nuclear migration. Embryo appears as a mass of yolk spherules (~0-16h). B, Nuclear migration (16-26h). C, Cumulus formation; contraction of blastoderm (26-40h). D, Cumulus migration begins; germ disc apparent (40-45h). E, Cumulus migration continues (46-52h). F, Cumulus migration ends (52h- 63h). G. Caudal bud forms; dorsal field begins to form(63h-72h). H, Caudal bud complete; cumulus disappates; dorsal field expansion (72h). I, Caudal bud migration, dorsal field expansion continues (72h-101h). J, Germ band formation, segmentation apparent (101-111h). K, Appendage bud formation (111-137h). L, Inversion begins: ventral sulcus appears along ventral midline; appendage buds elongate (137h-172h)). M, Mid-inversion (172h-205h). N, Inversion complete, ventral closure begins (205h-233h). O, Ventral closure complete (233h). Magnification x54, embryos are approximately 700 µm in diameter. Modified from Chaw, et al. 2007. 1 The first figure describes development in Achaearanea tepidariorum through germ band formation and segmentation. The second figure describes development in Zygiella x-notata. The Z.x-notata figure illustrates species differences and provides an idea of how post-germband development looks in A. tepidariorum.
0-10h 10-15h 15-25h 25-30h
30-40h 40-45h 45-55 55-65
Stages of early embryogenesis of the spider, Achaearanea tepidariorum, at 25º. Asterisks indicate the corresponding site of the different stage embryos. White areas in the illustrations indicate yolk. Scale bar: 200 µm. Modified from Akiyama-Oda & Oda, 2001.
2 Lepidopteran (Butterfly) Wing Development (April Dinwiddie)
Painted Lady (Vanessa cardui)
The Painted Lady (Vanessa cardui) is found throughout the world. It is also known as the Thistle butterfly, and the Cosmopolitan. As an adult, V. cardui is mostly black, brown, and orange with some white spots; the underside is gray with white and red markings.
Vanessa cardui Life Cycle:
Egg: Eggs are pale green, and hatch about 3 days after fertilization.
Larval Stages: Like other holometabolous insects, Lepidoptera go through complete metamorphosis. The larvae molt as they grow (each shedding represents a new “instar”, and there are five instars total).
Pupation: At the end of the fifth instar, the caterpillar will pupate. The chrysalis becomes almost transparent when the butterfly is about to emerge.
Adult: An adult will emerge about 7 to 10 days after the chrysalis has formed.
Wing disks develop in association with a trachea that runs along the base of the wing and are surrounded by a thin peripodial membrane, which is linked to the outer epidermis of the larva by a tiny duct. The wing disks are very small until the last larval instar, at which point, they increase dramatically in size, and are invaded by branching tracheae from the wing base (these branches precede the morphological formation of the wing veins). In the pupa, the wing forms a structure that becomes compressed from top to bottom and pleated from proximal to distal ends as it grows, so that it can rapidly be unfolded to its full adult size. Several patterns seen in the color of the adult wing are preceded by the expression of particular transcription factors (e.g. Engrailed, Spalt, Distalless, etc) in the late 5th instar and early pupa.
Forewing (L) and hindwing (R) discs of late forth/early fifth instar. Note milky, refractant (shiny), white trachea around base. (Anterior, top; posterior, bottom.)
Dissecting Lepidopteran Wing Disks
Larval Wings Disks (Butterfly) OPTIONAL: Anesthetize 4th or 5th instar larvae: place one larva at -20°C for 5 minutes (**Be careful not to freeze it!**) or multiple larvae at 4°C for 15-30 minutes (or longer).
Dissect out wing discs by one of the following methods:
Method A In a Sylgard coated dish in 1X PBS orient the larva so that dorsal is up (towards you) and ventral is down (towards the Sylgard). Put a pin through the larva around 3/4 of the way towards the posterior of the animal. While stretching the larva out, put another pin just behind the head carapace of the larva. (If the animal is not stretched out completely between the two pins, reorient pins/larva). • Using a pair of dissecting scissors, slice the skin of the larva from about the fifth segment to the base of the head. This slice should be along the dorsal midline of the animal. Be careful not to cut the underlying gut! • Carefully pull back the skin on one side; pin down if desired. The imaginal wing discs are located in the 2nd and 3rd thoracic segments, so you may want to pull/pin the skin at the 1st and 4th segments. (Hint: Use bristles to count segments; each segment has one D/V line of bristles.) You may want to gently push/pin aside the gut as well (do not put a pin thought the gut; brace the gut against a pin.) • Locate wing discs: They are a bit posterior and ventral to the lateral bristle, near the base of the larval legs in T2 and T3. The discs are somewhat transparent at younger stages (yellow/red at older stages), so you may want to look for the milky, refractant, white trachea that are attached to the base of each disc.
Method B In a Sylgard coated dish in 1X PBS orient the larva ON ITS SIDE. Pin and stretch the larva as described in Method A. • Use your forceps to tear the skin (make sure you do not tear deeper than the sink) behind the lateral bristle of the 2nd and 3rd thoracic segments. The imaginal discs will either “pop” out or will be sitting in the nearby tissue. Be careful not to harm the discs when you tear the skin
Fixation and Staining 1. Dissect out wing discs: carefully tear away the tissue at the base of each disc. Place in 1X PBS until all four wing discs are found.
2. Place discs in 3.2% PFA in PBS (you will be performing flourescent stains on butterfly discs) for 15-20 minutes. While fixing, dissect off remaining trachea and/or tissue from the discs. You can also try to remove the peripodial membrane, although antibody staining works with membranes on (in situs do not).
2 3. Wash in PT until ready to stain. Follow general Drosophila protocol for antibody stains (but consider leaving the disks in secondary overnight if they are large). Mount as for Drosophila disks, but consider using thicker coverslip or stack of coverslips for the side supports.
Pupal Wing Disks (Butterfly)
Collect pupae that are at least 12 hours old. Before this time, the cuticle is too soft and the dissection is technically very difficult, because it is hard to separate the cuticle from the forewing still attached to it.
OPTIONAL: Anesthetize pupae by placing them at -20°C for 5 minutes or at 4°C for 15-30 minutes (or longer).
1. Pin the pupa through the abdomen and thorax laterally so that the wing is accessible. 2. Cut the cuticle around the wing margin (forewing shape is visible on each side of the pupa). 3. Lift the cuticle and cut through the trachea that connects the forewing to the thorax. 4. Using fine forceps, remove the peripodial membrane that covers the hindwing. 5. Cut the trachea that connects the hindwing to the thorax
Place the wing discs in 3.2% PFA in PBS for about 20 minutes, wash in PT, and antibody stain as you would Drosophila embryos or disks.
3 Grasshoppers
1. Fill a Sylgard coated dish with 1X PBS (if staging ‘hoppers or fixing stages older than 30%) OR PEM-FA (if fixing stages younger than 30%—fix helps to hold delicate embryos together). 2. Place 1-3 grasshopper pods in dish. 3. Sink, clean off and orient the pods in the same direction. The “cap” is the slightly darker, granular area at one end of the pod (on right in diagrams). 4. Poke a hole in the side of each pod on the end opposite the “cap.”
5. With your forceps GENTLY squeeze the tip of the cap. This pushes the embryo away from the cap and you will see yolk stream out of the hole you made. 6. Cut the tip of the cap off with scissors and then use forceps to gently squeeze the middle of the pod. Squeeze until the entire embryo (they’re super long!) is out. The embryo sits on yolk, so squeeze out a lot of yolk as proxy for the embryo.
7. If embryo(s) are in 1XPBS, transfer them into a new Sylgard coated dish with PEM-FA and fix for 12-15 minutes. If embryos are already in fix, fix for 15 minutes (start fix time when you first poke a hole in the pod). 8. While the embryo is fixing, remove the amnion and any remaining yolk. If you are preparing for neural staining, make sure to rip open the membrane across the dorsal side of the embryo within the first few minutes of fixation. 9. Wash in PT until ready to start staining. 10. Separate grasshopper embryos into whatever number of eppendorf tubes you need to do your staining. Add a small number (50 is plenty) of rehydrated Drosophila embryos to each tube as internal controls. 11. Incubate in 300 µl PT+5% NGS and carry out antibody staining just as you would for Drosophila. For embryos 25-35%, you may want to extend the time in secondary antibody to 4 hrs, and for those older than 35%, you may want to leave in secondary overnight at 4°C (1:300 RT 2 hrs, then dilute to 1:600 4°C o/n). If your stains have a lot of background, use the secondary at 1:600. 3
Live grasshopper embryos shown at 5% developmental time intervals from 25% to 50% of embryogenesis. 5% development = 24 hours at 32°C
Appearance of metathoracic limb of live embryos at 5% developmental intervals from 25% to 60% of embryogenesis. Note that there are marked differences between each stage, particularly involving segmentation, flexion, invagination of apodemes and differentiation of musculature.
Adapted from Bentley et al., 1979
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Pax3/7 and Engrailed expression in developing grasshopper embryos (from Davis et al., 2001)
5 Crickets and Milkweed Bugs
Similar to grasshoppers, but you can briefly dechorionate (less than 1 minute; or skip bleach dechorionation altogether for crickets as you can see into egg easily).
Dissect embryo out into PEM-FA fixative. For younger stages, poke a few holes first to allow fix to enter for 1-2 minutes before dissecting. Fix for 15-20 minutes.
Wash in PT and proceed with standard protocol.
Embryonic Stages of the Common House Cricket (Acheta domesticus)
Development of the cricket Acheta domesticus, an intermediate germ band insect. (A) Major features of development. (1) The ventral germ band becomes immersed inside the yolk and migrates posteriorly. After establishing segmentation, the embryo revolves around the posterior end of the egg and assumes its final position. (2-4) The head emerges from the yolk and moves anteriorly. The embryo begins to progressively cover the yolk sac as tissues differentiate. (5, 6) The embryo closes its dorsal flanks. The leading edge contains the mesodermal cardioblasts. (7) Frontal view of the cricket embryo during the completion of organogenesis. (B) Schematic diagram of the development of the amniotic cavity in intermediate germ band insects. (1) Lateral folds starting at the edges of the germ band grow over the germ band. (2) The lateral folds meet ventrally beneath the germ band. (3) The membranes fuse, causing the separation of the amnion and serosa, thereby surrounding the embryo with yolk. (After Schwalm, 1997).
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Photomicrographs of embryonic stages of the house cricket Acheta domesticus (bars: 1 millimetre). a – zygote, b – start of embryonic development with formation of the germ band, c – mid-stage embryo with segmented body and development of thoracic/abdominal extremities, d – nearly fully developed embryo immediately before the dorsal closure, e – fully differentiated embryo before hatching, f – hatched larva of Acheta domesticus (Figure taken from Robert Sturm).
7 Milkweed bugs
8 Nipam H. Patel
Tribolium Whole Mount Procedure
1. Collect embryos. Place adults in flour containing no yeast, or flour containing 1% very finely ground yeast that is small enough to pass through the #50 sieve. After allowing the beetles to lay eggs for 1-3 days, sieve the adults away from the flour using a #25 sieve (Fisher Scientific Cat# 04-881N). Then sieve the eggs away from the flour using a #50 sieve (Fisher Scientific Cat# 04-881T).
2. Dechorionate the eggs with 50% bleach. Pipette bleach solution up and down and/or gently agitate the container to ensure all embryos are in contact with the bleach solution. When all the chorions and most of the flour particles are dissolved away, transfer the embryos to a fresh solution of 50% bleach for another one or two minutes. The total time required in the bleach will vary; more time is needed if there are lots of embryos and/or lots of flour. We usually keep the eggs in bleach for anywhere between four and ten minutes. To little time in the bleach will leave traces of the chorion on the embryos and reduce the efficiency of the following fixation and devitellinization steps. Too much time in the bleach will cause the disintegration of the embryos, particularly those that are at the blastoderm stage or younger.
3. Rinse well with dH2O.
4. Place embryos into a glass scintillation vial containing:
Heptane : PEM-FA in a ratio of 1:1 (v:v)
PEM 0.1M Pipes 2mM EGTA 1mM MgSO4 Adjust pH to 6.95 with conc HCl Can be stored at 4°C
PEM-FA 9 parts PEM to 1 part 37% formaldehyde (Fisher, Cat # F79-500) Make just before use.
I usually use scintillation vials that hold about 20 ml total, thus I use 10 ml heptane, 9 ml PEM and 1 ml 37% formaldehyde. Try not to put in more embryos than will fill a monolayer at the heptane/fix interface. You can gently agitate the embryos every now and then while they are fixing, but vigorous shaking is unnecessary. Fix the embryos for 15 minutes in this manner. Overfixation will reduce staining intensity for many antibodies, underfixation will result in poor morphology and high background with some antibodies.
5. Remove the aqueous layer (PEM-FA, lower phase) with a pipette. Remove as much of the PEM-FA as possible. Then remove most of the heptane, but don’t let the embryos dry out. I try to remove all but the last 500µl or so of the heptane.
1 Nipam H. Patel
6. Add about 9 ml of –40 to –70°C heptane and then place the scintallation vial at –70°C for 5 minutes. Then add an equal volume of room temperature methanol. Warm under running tap water to room temperature while shaking vigorously by hand for about 20- 30 seconds. The step of cooling to –40 to –70°C slightly improves the devitellinization efficiency, but is not absolutely necessary.
7. Those embryos which are devitellinized will fall to the bottom and will predominately consist of stages prior to germband retraction. Those embryos which are not devitellinized will stay at the interface and are mostly embryos that have completed germband retraction. Both sets of embryos are usable for antibody staining. The embryos at the bottom (“cracked”) should be transferred with a pipette to a new tube of absolute MeOH. Those at the interface (“uncracked”) can be recovered by first removing as much heptane (top phase) as possible from the scintillation vial. Then remove most of the MeOH. Now refill the vial with MeOH. The uncracked embryos should now sink to the bottom since there is no longer a heptane phase for them to sit in. These uncracked embryos can now be transferred to another tube of absolute MeOH. They can still be used for antibody staining because virtually all these embryos do have slits in the vitelline membrane that will allow the antibodies access to the embryos. I usually keep the two types of embryos, cracked and uncracked, separate throughout the remaining procedures.
8. Wash the embryos five times with absolute MeOH to remove residual heptane. At this point, the embryos can be stored in absolute MeOH at –20oC for several months with only a slight decrease in their staining intensity for most antibodies I have tested. As needed, aliquots of embryos can be removed and processed as described below.
9. Remove MeOH and replace with PBT. I carry out all the subsequent steps in 5 ml snap cap polypropylene tubes (Falcon # 2063). PT: 1x PBS 0.1% Triton X-100
10. Immunostain as you would Drosophila
2 Nipam H. Patel
Tribolium (Flour beetles)
Similar protocol to Drosophila dechorionization and immunohistochemistry.
Figure 1. Relationships of insect orders, germ types, and ovary types. a, Phylogenetic tree of insect orders. Germ-type: S, short; I, intermediate; L, long. Ovary type: Pan, panoistic, no nurse cells; Mer, meroistic, with nurse cells. Germ types do not form separate monophyletic groups, and certain orders include representatives of all three germ types. One general trend is visible: long-germ development appears to be restricted to the more phylogenetically derived orders, which also contain meroistic ovaries. b-d, DAPI-stained ovaries. o, Ooctyes; f, follicle cells; g, germarium. b, The panoistic ovary of the Orthopteran Schistocerca americana. c, The meroistic ovary of the Hemipteran Oncopeltus fasciatus. d, The meroistic ovary of the Dipteran, Lucillia cuprina.
3 Nipam H. Patel
Figure 2. Tribolium even-skipped protein expression at different stages of development. Nomarski (a,c,d,f,g, and i) and fluorescence (b,e, and h) images of embryos stained with mAb 2B8 and counterstained with DAPI. Triangles and arrows mark the anterior boundaries of even-skipped primary stripes 1 and 2, respectively. Arrowhead marks the position of even-skipped primary stripe 3.
Figure 3. Relationship of even-skipped (brown) and engrailed (black) expression in Tribolium. Protein detected using an HRP histochemical secondary. The inclusion of nickel chloride when developing the engrailed secondary results in a black reaction product. Engrailed was detected with 4D9 and even-skipped with 2B8.
4 Nipam H. Patel
Figure 4. Comparison of even-skipped and engrailed patterns in Schistocerca and Tribolium uring abdominal segmentation. Tribolium (Tri; a and c) and Schistocerca (Sch; b and d) embryos stained for engrailed in black and even- skipped in brown (a and b) or even-skipped in black and engrailed in brown (c and d). In all four panels, a diamond marks engrailed stripe 5 (second thoracic segment). The Tribolium and Schistocerca in a and b are at similar stages of generating segments because both have six engrailed stripes. Likewise, embryos in c and d are at similar stages because both have just started to form the ninth engrailed stripe.
Stem cells in the germarium produce oocytes that are not attached to nurse cells. The germinal vesicle of the oocyte transcribes all of the maternal RNA of the egg. The small, flat cells surrounding the oocyte are the follicle cells a and b, Initial even-skipped expression in the posterior half of the embryo. c, The first even-skipped primary stripe forms as even-skipped protein disappears from a circumferential interstripe zone. An irregular “wave” of mitosis the begins and even-skipped protein temporarily diffuses from the nucleus to the cytoplasm of mitotic cells (not shown). d and e, The primitive pit begins to form and cells continue to aggregate and divide to form the embryonic germ anlage. Gastrulation now begins at the ventral midline. The second even-skipped primary stripe forms anterior to the primitive pit.
5 Nipam H. Patel
Mosquito (Anopheles gambiae) Whole Mount Procedure
1. Use water to rinse embryos into a very fine Nitex mesh (20 - 50 µm opening). The Nitex mesh usually used in my lab to hold Drosophila embryos (85 - 100 µm openings) will let some of the mosquito embryos slip through as the Anopheles embryos are much narrower than Drosophila embryos.
2. Dechorionate the eggs with 50% bleach. Pipette bleach solution up and down and/or gently agitate the container to ensure all embryos are in contact with the bleach solution. The total time required in the bleach will vary with the stage of development and the particular mosquito species being used. I usually keep the Anopheles eggs in bleach for anywhere between one and four minutes. Too little time in the bleach will reduce the efficiency of the following fixation and devitellinization steps. Too much time in the bleach will cause the disintegration of the embryos. The younger stages (before gastrulation) seem particularly sensitive to the amount of time they are in bleach. This bleach treatment removes the thin, waxy chorion, but there will still remain a black endochorion, but it is still possible to antibody stain and look at the embryos without removing it (see below).
3. Rinse well with dH2O to remove bleach.
4. Place embryos into a glass scintillation vial containing:
10 ml heptane 9 ml PEM 1 ml 37% formaldehyde
PEM 100 mM Pipes (Disodium salt, Sigma Cat. No. P-3768) 2 mM EGTA 1 mM MgSO4 Adjust pH to 6.95 with conc HCl Can be stored at 4°C
37% formaldehyde (Fisher, Cat # F79-500)
Try not to put in more embryos than will fill a monolayer at the heptane/fix interface. You can gently agitate the embryos every now and then while they are fixing, but vigorous shaking is unnecessary. Fix the embryos for 15-20 minutes in this manner. Overfixation will reduce staining intensity for many antibodies, underfixation will result in poor morphology and high background with some antibodies.
5. Remove the aqueous layer (PEM-Formaldehyde; lower phase) with a pipette, eliminating as much of this aqueous layer as possible. Then remove most of the heptane, but don’t let the embryos dry out. I try to remove all but the last 500µl or so of the heptane.
1 Nipam H. Patel
6. Add about 9 ml of –40 to –70°C heptane and then place the scintillation vial at –70°C for 5 minutes. Then add about 9 ml of room temperature methanol. Warm under running tap water to room temperature while shaking vigorously by hand for about 20- 30 seconds. Do not worry that some embryos stay at at the heptane/methanol interface and others are at the bottom. The step of cooling to –40 to –70°C slightly improves the "cracking" efficiency, but is not absolutely necessary.
7. Remove as much of the heptane as possible and most of the methanol; leaving behind a little methanol to keep the embryos from drying. Most all the embryos, even those that stayed at the interface, will have small cracks in the endochorion and vitelline membrane which will allow antibody penetration.
8. Wash the embryos five times with absolute MeOH to remove residual heptane. If embryos have black pigmentation in the endochorion (as is the case with Anopheles gambiae), this coloration will need to be lightened. This can be done by placing the embryos into a freshly prepared solution of 67% methanol/32% water/1% hydrogen peroxide for 24 hrs at room temperature with gentle mixing. I use about 5 ml of this "bleaching" solution for every 100 µl of packed embryos. After 24 hrs, the black coloration is changed to an amber color which is light enough to see staining through (particularly if the final DAB reaction is nickel intensified to yield a black product). This hydrogen peroxide "bleaching" step can be extended for up to three days if needed (replacing and adding fresh solution each day), but extending the "bleaching" past 24 hrs does lower antibody staining intensity somewhat. [Note: recently I have had good luck with bleaching salamander embryos in a solution of 80% methanol/10% water/and 10% hydrogen peroxide for 12 hrs. This might work also well for the mosquites, but I have not tried it on them yet.]. After "bleaching", wash the embryos extensively with absolute methanol. At this point, the embryos can be stored in absolute MeOH at –20°C for several months with only a slight decrease in their staining intensity for most antibodies I have tested. As needed, aliquots of embryos can be removed and processed as described below.
9. Remove MeOH and replace with PT. I carry out all the subsequent steps in 5 ml snap cap polypropylene tubes (Falcon # 2063).
PT: 1x PBS 0.1% Triton X-100
10. Continue immonostaining as you woul Drosophila
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