Arthropod/Fly Lab 2010 June 21-26

Lectures: Celeste Berg Iswar Hariharan Nipam H. Patel Matthew Ronshaugen

Labs: Bill Browne Elise Delagnes April Dinwiddie Abby Gerhold Yassi Hafezi Hannah Rollins TABLE OF CONTENTS

I. INTRODUCTION 2 II. SCHEDULE 3 III. EXPERIMENTAL OVERVIEW 4 IV. PROTOCOLS 13 IV.1 General Fixation and Antibody Staining 13 IV.2 Rapid Antibody Staining Protocol 15 IV.3 Histochemical Development Reactions 16 IV.4 Labeling with Multiple Primary Antibodies 18 IV.5 Fixing and Staining Other Arthropods 19 A. Spider 19 B. Artemia 20 C. Mysids 21 D. Grasshoppers 22 E. Parhyale 23 IV.6 Fixing and Staining Post-embryonic Tissues (Imaginal Discs) 27 A. Drosophila Wing Imaginal Discs 27 B. Butterfly Wing Imaginal Discs 29 IV.8 In situ Hybridization 30 IV.9 Injection in Parhyale 33 V. GENERAL SOLUTIONS 36 V.1 Fixatives 36 V.2 Solutions for antibody protocols 36 V.3 Solutions for in situ hybridization 38 VI. MAKING DISSECTIONS TOOLS 40 VI.1 Making blunt probes 40 VI.2 Making Tungsten needles for dissecting 40 VII. AVAILABLE STOCKS & REAGENTS 41 VII.1 Antibodies Separate Handout VII.2 In situ Probes 42 VII.3 Fixed Embryos 43 VII.4 Drosophila Stocks 44 VIII. DEVELOPMENTAL BIOLOGY & ANIMAL STAGING 46 VIII.1 Parhyale 46 VIII.2 Drosophila 51 VIII.3 Spiders 65 VIII.4 Grasshoppers 67

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 and analyze mutant phenotypes. In order to do that, you will be performing different molecular and embryological techniques, such as antibody staining, in situ hybridization, live imaging, and lineage tracing. At first and most importantly, we would like you to use a set of antibodies to detect the expression of important developmental proteins in the fruit fly Drosophila melanogaster. This step will allow you to master the procedure of antibody staining while studying the spatiotemporal expression of these proteins. Because these proteins are strongly conserved in all arthropods studied to date, this set of 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. You can stain these tissues with available antibodies and compare the expression pattern of various proteins between flies and butterflies. We will also conduct a clonal analysis experiment in which we will ask whether certain mutations affect the sorting, positional or adhesive properties of cells with the developing wing disc. We 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.

2 II. Schedule

Monday (lecture: Intro to Embryogenesis and Post-embryonic Development of Drosophila) Look at pre-stained embryos Rapid staining Review of available antibodies, in situ probes, and fly stocks Dissection of imaginal disks Imaging 101A

Tuesday (lecture: Growth Control) Parhyale injections Imaginal disk cell sorting experiment Other arthropods Chalk Talk

Wednesday (lecture: Arthropod Evo-Devo) More arthropods (plankton tow) Ovary dissection Imaging 101B

Thursday (lecture: Non-coding RNAs) Continue experiments Snorkeling (weather permitting)

Friday (lecture: Oogenesis) Continue experiments

Saturday (lecture: Butterfly wing patterning) Prepare for Show N’ Tell Show N’ Tell

Sunday Whaling

3 III. Experimental Overview

III.1 Observations of Embryogenesis A. Antibody Staining of Drosophila Embryos Antibody staining Drosophila will prepare you for staining other arthropods. In this experiment, you will investigate protein expression patterns throughout Drosophila development in the following gene classes (appropriate antibody in parentheses): pair-rule (DP312), segment polarity (DP312), Hox (FP3.3), and axons/neurons (BP102, 9F8, DP312). 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 along the length of each bench. Each group should complete all six of the following antibody stains on Drosophila. The rapid antibody protocol is in section IV (on page 15) of this manual.

** DO NOT USE ALL OF YOUR FLIES. These flies are a mix of embryos ranging from 0-16 hours. Only use 15µl of settled fly embryos in MeOH per 1.5ml eppendorf tube. This will be about 20µl when rehydrated. If you are unsure how many flies to use, ask for help. We will have examples showing a good amount of embryos to use. Using to many or too few embryos can effect the signal to noise ratio of your staining.

React Antibody Staining Pattern µl:300 1:300 with Perform the following stains as rapid antibody stains (1-day) Anti-Pax3/7 (pair-rule, segment polarity, neural DAB+ 1) DP312 gene family) In Drosophila, pair-rule pattern 10.0 Ni early, then segmental pattern, then CNS pattern Anti-Ubx (homeotic gene). In Drosophila, DAB+ 2) FP3.3 15.0 regional expression pattern Goat anti-mouse Ni HRP conjugated DAB+ 3) BP102 Stains all CNS axons (unknown antigen) 5 Ni 4) BP102 See above 5 AEC Stains nuclei of all neurons (elav gene product, DAB+ 5) 9F8 15.0 encodes neural-specific splicing factor) Ni Goat anti-mouse AP BCIP/ 6) 9F8 See above 15.0 (115-055-166) NBT

4

B. Into the Wild – Comparative Antibody Staining in Other Arthropods

Use the protocol and expertise you have acquired from your Drosophila antibody stains to detect the expression of these developmental proteins in many different arthropod species. Besides the additional arthropods listed below, you can collect specimens from the area around Woods Hole. Enjoy!

Insects: Junonia (Precis) coenia (buckeye butterfly) Tribolium castaneum (flour beetle) Schistocerca americana (grasshopper) Apis millifera (European honeybee)

Crustaceans: Parhyale hawaiensis (amphipod - beach hopper) Chelura sp. (amphipod - beach hopper) Caprella sp. (amphipod – skeleton shrimp) Limnoria sp. (isopod) Cyathura sp. (isopod) Libinia sp. (spider crab) Pagurus longicarpus (long-claw hermit crab) Emerita talpoida (mole crabs) Crangon septemspinosa (sand shrimp) Palaemonetes sp. (grass shrimp) Mysidium columbiae (opossum shrimp) Balanus sp. (barnacle) Artemia salina (brine shrimp - “sea monkeys”)

Chelicerata: Parasteatoda tepidariorum (common house spider) Limulus polyphemus (horseshoe crab) Tanystylum sp. (pycnogonid - sea spider) Callipallene sp. (pycnogonid - sea spider)

Some suggestions: 1) Examine the expression of Engrailed, Pax3/7, FP6.87, and the pattern of axonogenesis with αHRP at various stages in an arthropod of your choice. 2) Do a double antibody stain with Pax3/7 plus Engrailed and/or Ubx/abdA plus Pax3/7 in Parhyale or any other arthropod of your choice. What is the pattern and how does it compare to Drosophila? In Parhyale, can you identify the different rows in which each gene is expressed?

5 C. 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). Stains to do and compare: 1) 1G10 (anti-Hunchback) on Drosophila 2) 7C11 (anti-Hunchback) on Grasshoppers (15 - 25%) 3) 2B8 (anti-Eve) on Drosophila, Tribolium and Mysids stage 1 and 2 (sonicate stage 2) 4) 3B9 (anti-Eve) on Grasshopper (15-25%) 5) 7H5 (anti-Eve) on Artemia (sonicated) 6) RαPhEve (anti-Eve, ask Nipam for antibody) on Parhyale 7) Double label 2B8 (Black, the rapid protocol in the protocol section will work on this) + 1G10 (Brown) on Drosophila 8) Double label 7C11 (Black) + DP312 (Brown) on Grasshoppers (15-25%) 9) Double label 2B8 (Black) + 4D9 (Brown) on Tribolium

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? Stains to do and compare: 1) DP312 (anti-Pax 3/7) on Drosophila, Tribolium, Oncopeltus, Grasshoppers (15 – 30%), Parhyale, Mysids, stage 1, 2, and 3 (sonicate stage 2 and 3), Artemia (sonicated), Spiders, and Limulus 2) 2B8 (anti-Eve) on Drosophila, Tribolium, Mysids stage 1 and 2 (sonicate 2) 3) 3B9 (anti-Eve) on Grasshopper (15-25%) 4) 7H5 (anti-Eve) on Artemia (sonicated) 5) RαPhEve (anti-Eve) on Parhyale 6) 4D9 (anti-engrailed) on Drosophila, Grasshoppers (15 – 30%), Tribolium, Parhyale, Mysids, stage 1,2, and 3 (sonicate stage 2 and 3) 7) 4F11 (anti-engrailed) on Artemia (sonicated) 8) Double label DP312 (Black) + 4D9 (Brown) on Drosophila, Tribolium, Grasshoppers, and Parhyale 9) Double label DP312 (Black) + 4D9 (Brown) on Mysids stage 1 and 2 (sonicate stage 2) 10) Double label DP312 (Black) + 4F11 (Brown) on Artemia (sonicated)

6 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. Stains to do and compare: 1) FP6.87 (anti-Ubx/abdA) on Drosophila 2) FP6.87(anti-Ubx/abdA) on Grasshopper (25 – 40%) 3) FP6.87 (anti-Ubx/abdA) on Artemia (sonicated) and Mysids (sonicated Stage 2 and 3) 4) FP6.87 (anti-Ubx/abdA) on Parhyale and Spiders 5) 8C11 (anti-Antp) on Drosophila 6) 1D11 (anti-Scr) on Drosophila

Neurogenesis and Axonogensis Examine the process of Drosophila neurogenesis and axonogenesis. Compare neurogenesis and axonogenesis in Drosophila to neurogenesis in other arthropods Stains to do and compare: 1) 1G10 (anti-Hunchback) on Drosophila 2) 7C11 (anti-Hunchback) on Grasshoppers (20-40%) 3) 2B8 (anti-Eve) on Drosophila 4) 3B9 (anti-Eve) on Grasshopper (30-40%) 5) RαPhEve (anti-Eve) on Parhyale 6) 1D4 (anti-FasII) on Drosophila 7) 8C6 (anti-FasII) on Grasshopper (35-45%) 8) 3B11 (anti-FasI) and 6F8 (anti-FasIV) on Grasshopper (35-45%) 9) 22C10 (anti-CNS/PNS) on Drosophila 10) Goat anti-HRP on Artemia and stage 3 Mysids (all sonicated) 11) Goat anti-HRP on Drosophila 12) Goat anti-HRP on Grasshopper) 13) Goat anti-HRP on Parhyale and Spiders 14) Double label 9F8 (Black, this works with the rapid staining protocol in the protocol section) + 22C10 (Brown) on Drosophila 15) Double label 3B9 (Black) + 7C11 (Brown) on Grasshoppers (30-40%) 16) Double label 2B8 (Black) + 22C10 (Brown) on Drosophila

Neurogenesis and Axonigenesis Fluorescently Triple label the fly CNS with antibodies made in different animal species (mouse, rabbit, rat). Animal Antibody Secondary & Notes Mouse DP312 (anti-Pax3/7+) GαM 555 HIGHLY CROSS ABSORBED OR 4D9 (anti-engrailed) GαM 555 HIGHLY CROSS ABSORBED Rabbit 10900 (anti-eve) GαRb 647 CONFOCAL ONLY Rat 7E8 (nuclei of all neurons) GαR 488 HIGHLY CROSS ABSORBED None anti-HRP FITC (all neurons) (no secondary needed)

Additional Genes We will provide you with a list of additional antibodies (Additional antibodies printout) you can add to your experiments. 7

D. Live Imaging of Embryos Collect Drosophila embryos at the right stage (as close to fertilization as possible if you want to look at early nuclear divisions) and mount on glass slides or biofoil, covered with halocarbon oil and a cover slip. Try live imaging the following: 1) Wild type 2) GFP lines (e.g. moesin:GFP, histone:GFP…) 3) GAL4 lines (e.g. Dpp-Gal4, wg-Gal4) crossed with UAS:GFP or G-TRACE 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 would like you to try injecting Parhyale (we will give demonstrations). First, try injecting a one-cell embryo, then try moving up to two- through eight-cell embryos (see the electronic version of the protocol for more suggestions). After you have successfully injected some embryos, we would like you to try filming them. Alternatively, and/or if your injected embryos die, you can make a movie of Parhyale development with one of the transgenic lines. Reagents available for Parhyale live imaging: Labeling Reagent/ Usage Information Antibody/ Notes Transgenic Line (Live Embryos) Fixed Embryos NLS-DsRed mRNA Stock ~1ug/ul n/a Nuclear-localized red fluorescent Use b/w 500ng/ul to protein- will label all progeny of 1ug/ul injected cell. Will fluoresce brightly ~1 day after injecting FITC-Dextran 50mg/ml stock Anti-Flourescein Green fluorescent dye- able to (Fluorescein isothiocyanate- Use at 10-20mg/ml for Use at final conc. of visualize immediately dextran) tracing, and >25mg/ml 1:3000 -too much will kill cell almost for ablations immediately -to ablate cell lineage later in development, shine blue light on FITC+ cells TRITC-Dextran 50mg/ml stock n/a Red fluorescent dye- able to (Tetramethylrhodamine Use at 25mg/ml-50mg/ml visualize immediately isothiocyanate–dextran) for lineage tracing

HS-NLS-DsRed Parhyale Heat shock embryos: n/a Nuclear-localized red protein- after (transgenic line produced by 1 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)

8 E. 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. 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 Look at ventral furrow formation using DEcadherin-GFP Observe early nuclear division cycles using GFP-nls 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.

F. 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. Try α- Engrailed, α−wingless, α−eve and α−AbdB on the Kruppel mutant embryos. Additionally the cuticles of this mutant will show some interesting defects.

Cuticle preps of various gap, homeotic and segmentation gene mutants: Refer to Nusslein-Volhard and Wieschaus (1980 Nature) for background and significance. Cuticle preps are an excellent system in which to study various signal transduction pathways and to establish epistasis. You can do these in Drosophila 1) Analyze the denticle belt patterns for embryonic patterning mutants: e.g. Kr, eve, ftz, wg, en, ptc, hh. 2) You can also collect embryos from these crosses and stain them with various antibodies.

G. Parhyale RNAi You can look at the finction of the single-minded gene in Parhyale using sim siRNA injection.

9 H. 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 back of manual for a 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! Think 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) Tribolium castaneum (flour beetle) embryos are great for fluorescent work. There is a list of possible ISH probes in the back of the manual. Apis embryos tend to be difficult to work with and fail to stain evenly, but occasionally make beautiful pictures. Caveat emptor! 3) 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 4) 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. 5) 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. 6) Observe the differential expression of the related but differentially expressed genes SoxN and Dichaete in the nervous system. 7) Compare the expression pattern of midline patterning genes (e.g. sim) between Drosophila and Apis embryos.

10 III.2 Larval Structures & Imaginal Disc Development

A. Wing Imaginal Disc Staining Look at the expression of Engrailed, Spalt and Ubx orthologs via antibody staining in Drosophila and in the buckeye butterfly. Engrailed marks the posterior wing compartment (and, in butterflies, the center of the eyespot), Spalt is a readout of dpp signaling (proven in Drosophila) and will mark butterfly eyespots, and Ubx marks the hindwing. Stains to do and compare: 1) 4F11 (anti-Engrailed) on Drosophila wing discs (3rd instar) 2) 4F11 (anti-Engrailed) on buckeye wing discs (4th and 5th instars) 3) FP6.87 (anti-Ubx) on Drosophila wing discs (3rd instar) 4) FP6.87 (anti-Ubx) on buckeye 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) Use fluorescent secondaries on discs (while Drosophila discs will stain fine histochemically, butterfly wing disc staining works best fluorescently). Only late 5th instars and older butterfly larvae will have eyespot staining. For best eyespot stains, use discs from 12hrs post pupariation.

B. Clonal Analysis in the Drosophila Wing Imaginal Disc See Section VIII.2 for a description of how clones are generated in imaginal discs. In imaginal discs, there is very little cell migration. Thus the clone and twin spot are almost always immediately adjacent to each other. Their position with respect to each other is usually random. This has been studied extensively in the wing imaginal disc where clones tend to be elongated along the proximo- distal axis. It was noted by Garoia et al., that when clones mutant for the protocadherin fat are generated, that the mutant clones preferentially occupies the more proximal position. There are reasons for thinking that there may be gradients of cell affinity along the proximodistal axis, at least with respect to adhesion mediated by specific molecules (e.g. fat). Thus it is possible that cells that are mutant for fat adhere preferentially with cells that are found in the proximal wing (Similar gradients of cell affinity are thought to operate during vertebrate limb development). Some of the models that have been proposed to explain how an organ achieves its final size involve gradients of positional information. It would therefore be of interest to test whether a variety of mutations that are known to affect the growth and possibly the “sorting” properties of cells in the wing-disc are able to alter the adhesive properties of the cell with respect to the proximo-distal axis. We propose to generate clone/twin spot pairs for each of these mutations and assess their relative position along the proximo- distal axis of the wing disc. Students will gain familiarity with a number of important techniques used by Drosophila geneticists as well gain potentially novel insights into the relationship between growth and positional information in the developing wing. The wing imaginal disc is also compartmentalized (see section VIII.2) in non-mixing lineage restricted regions. While we know a fair amount about how these boundaries are established, the mechanisms that maintain these strict lineage restrictions are still not known. Try generating mutant clones within the wing discs and looking to see if any of these are able to “cross” the compartment boundary. You can mark the A/P boundary using an engrailed or cubitus interruptus (ci) antibody. We will provide a list of available reagents and stocks for various clonal analysis experiments.

11 III.3 Gonadogenesis and gametogenesis Embryonic, larval and adult ovaries can be stained with useful markers throughout development. 1) Gonadogenesis throughout development: Antibodies: Engrailed (terminal filaments), Spectrin (cytoskeletal component), Adducin (fusome component), Bic-D (oocyte specification)… 2) How is embryonic patterning established during oogenesis? For example… Stain ovaries with labeled Phalloidin to observe ring canals connecting nurse cells to the oocyte. Use live imaging to observe cytoplasmic and yolk flows in the developing oocyte.

12 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. 13 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 as noted in the table at the back of the manual. 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 for 30 minutes, and then replace with 300 µl 70% glycerol. AlexaFlour conjugates are very fade resistant even without the addition of an anti-fade compound. For HRP or alkaline phosphatase (AP) conjugated secondary antibodies, proceed to the histochemical development reactions (section IV.3)

14 IV.2 Rapid Antibody Staining Protocol

While this protocol produces antibody stains in one day, it only works well on very robust antibodies.

1. Rehydrate with 2X 1 min followed by 1X 10 min with PT.

2. Incubate 10 min in 100 µl PT+NGS.

3. Add primary antibody to the appropriate final concentration. Primary antibodies may be used at about 1.5 to 2 times the “normal” concentration (i.e., that used for the regular staining procedure).

4. Mix and incubate in the primary antibody at room temperature for 30 min.

5. Wash 3X 1 min with PT.

6. Wash 3X 10 min with PT.

7. Add secondary antibody. It is not necessary to pre-block with PT+NGS. For HRP immunohistochemistry, use the goat anti-mouse IgG at a dilution of 1:300. Mix and incubate in the secondary antibody for 30 min at room temperature.

8. Wash 3X 1 min with PT.

9. Wash 3X 10 min with PT.

10. React with the appropriate reaction protocol. (See general antibody prootocol, IV.1)

11. Wash with PT.

12. Start your next staining (if doing multiple labels). If you are done, put the embryos into 200µl 50% glycerol with DAPI for 10 – 30 min and then into 300 µl of 70% glycerol. The embryos will be fine in glycerol for several weeks at room temperature, several years at 4°C, and several decades at –20°C. Staining may fade (even over just a few hours) if your glycerol solution is acidic.

15 IV.3 Histochemical Development Reactions:

Unless specified, reagents are on FRONT SIDE BENCH or in the Beddington refrigerator/freezer

A. Black HRP Reaction (THIS IS THE DEFAULT REACTION) 1. Get DAB ready to add. DAB is a carcinogen and should be treated with great care. Frozen aliquots of 3 mg/ml DAB have already been made and are in the -20°C freezer. Thaw the 3 mg/ml DAB and dilute with 1X PBS and Tween-20 to make a DAB solution that is 0.3 mg/ml DAB and 0.05% Tween-20. Add 8 µl of 8% nickel chloride for each 1.0 ml of diluted DAB. Mix by inverting the tube several times. Treat all DAB waste with care. Liquid waste should go into the “liquid DAB waste” container in the hood of room 258. All solid waste (used tips and eppendorf tubes) should go into a zip-locked bag labeled “solid DAB waste” in the hood of room 258. Please do you best to minimize the amount of waste DAB that you generate (both liquid and solid). 2. After the last PT wash, drain the tubes down to about 1 mm above the embryos. Add 300 µl of the DAB+Ni solution. Make sure there are NO bubbles on the top of the tube. Mix very gently by spinning, again making sure that there are no bubbles on top. 3. Prepare a 0.3% solution of hydrogen peroxide by mixing 10 µl of 30% hydrogen peroxide with 900 µl of 1X PBS. This solution is only good for about 30 minutes. 4. Add 15 µl of the 0.3% hydrogen peroxide to each tube of embryos in DAB+Ni. Mix gently (by spinning), but quickly and thoroughly. Avoid introducing bubbles. If you do introduce bubbles, remove them from the surface. 5. Watch reaction down the dissecting scope. Leave the embryos in the tube and sight down with one eye (make sure to be illuminating from above with a white surface, such as a Kimwipe, below the tube). You can practice beforehand using one of your tubes of pre-stained embryos (drain down to 300 µl level of glycerol). 6. Let the reaction proceed until you have a good signal to noise ratio. The biggest mistake people make is stopping the reaction too soon. A little background is fine. Weak signal is a problem. You can safely let the reaction carry on for at least 5 – 10 minutes. 7. Stop the reaction by draining off the DAB (and put it into the DAB liquid waste container) and washing several times with PT (put the first wash into the liquid DAB waste, subsequent washes can go down the sink).

B. Brown HRP Reaction Follow the same protocol as above, but leave out the addition of nickel chloride.

C. Red HRP Reaction A red reaction product can be obtained using 3-Amino-9-ethylcarbazole (AEC; available from Sigma Cat. No. A-6926). 1. Wash embryos 1 X 5 min. in AEC buffer (300 µl). 2. Place embryos into 200 µl of the AEC substrate solution. 3. Add 15-20 µl of 0.3% hydrogen peroxide. Monitor reaction, although this can be difficult because the solution will turn cloudy. 4. After reaction is completed (10-15 min), wash 2 X 5 min PT. 16

D. Purple AP Reaction 1. Wash embryos 1 X 5 min in A.P. buffer (300 µl). 2. Add 200 µl of BCIP/NBT solution (pre-mix in separate eppendorf tube).

BCIP/NBT solution 1 ml A.P. Buffer 4.5 µl NBT (50 mg/ml in 70% DMF) 3.5 µl BCIP (50 mg/ml in 70% DMF) 3. Monitor the reaction. An optimal signal-to-noise ratio will usually be reached anywhere from 5-15 min, but the reaction can be allowed to continue for several hours if needed. The sensitivity of this technique is equal or better than a black HRP reaction. Alkaline phosphatase reactions, however, are more prone to background problems than HRP reactions and the reaction products sometimes give diffuse staining around fine structures such as individual axons. 4. Stop the reaction by washing 2 X 1 min with PT. Note for fly staining: Background alkaline phosphatase activity in cuticular stripes and in the trachea are sometimes observed in embryos past stage 17. Note that this background is not effectively inhibited by Levamisole, which is often used as an inhibitor of endogenous alkaline phosphatase in vertebrate tissues.

E. Blue AP Reaction 1. Follow steps 1-4 in the protocol for the purple alkaline phosphatase reaction above. 2. After stopping the reaction by washing with PT, wash the embryos 2 X 5 min and 1 X 30 min in methanol. The signal will slowly turn from purple to blue. Wash with methanol until the desired level of blue is obtained. The color change occurs because the purple alkaline phosphatase reaction product is actually composed of two different reaction products: an alcohol-soluble purple product and an alcohol-insoluble blue product. Because this procedure lowers the level of signal, it should be used only if the starting purple reaction product is relatively strong. 3. Wash 2 X 5 min PT After completing your histochemical reactions, either wash for an additional 20 min in PT and start your next staining (if doing multiple labels). If you are done, put the embryos into 200 µl 50% glycerol with DAPI for 10 – 30 min and then into 300 µl of 70% glycerol. The embryos will be fine in glycerol for several weeks at room temperature, several years at 4°C, and several decades at –20°C. Staining may fade (even over just a few hours) if your glycerol solution is acidic.

17 IV.4 Labeling with Multiple Primary Antibodies

A. Antibodies Made in Different Animals (Flourescent and Histochemical) If all your primary antibodies are made in different animals (for example: mouse, rabbit and/or rat), then you can do all of your stains simultaneously. For histochemical stains, this will only work if your secondaries are conjugated to different substrates (HRP and AP, but not both HRP or both AP). If your secondaries are conjugated to the same substrate, perform the color reactions sequentially. Make sure to let you reactions go to completion—at least 10 minutes. Wash well with PT between reactions!

B. Antibodies Made in the Same Animal (Histochemical) If your primary antibodies are made in the same animal (for example: mouse), then you will have to do your antibody stains sequentially. To cut down on the length of this process, you can do the first reaction following the rapid staining protocol, if your first antibody is particularly robust. This rapid protocol can then be followed by a regular length protocol. If all of your antibodies are “not-so-robust”, then follow a regular length protocol with another regular length protocol. Examples of robust antibodies are BP102, 22C10, and DP312. Others are noted in the suggested experiments, but feel free to ask about additional antibodies.

C. General Notes If you are using HRP-conjugated antibodies, make sure you do your darker reaction (black reaction product) first, followed by your lighter reaction (brown reaction product) and then your lightest reaction (red reaction product). If you are using two AP-conjugated antibodies, make your first reaction blue, and your second reaction purple (otherwise, both will be blue!). If you are using HRP-conjugate(s) together with AP-conjugate(s), you can either do the HRP reaction(s) first or the AP reaction(s) first. Wash with Glycine (pH=2) if your first reaction does not “stop”.

18 IV.5 Fixing and Staining Other Arthropods

A. Spider

Remove thee eggs from the egg sac before you start this protocol. Eggs develop approximately synchronously and so each brood can be staged by placing one in mineral oil. We have already done this for you. 1. 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** 2. Rinse. Transfer embryos to an eppendorf tube and rinse carefully with 1x PBS. Five quick washes should remove all the bleach. 3. 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.

19 B. 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.

20 C. Mysids

These embryos have already been fixed for you (fixed while still in the female brood pouch). They were stored in methanol, and have been rehydrated in PT (3x10min PT at room temp). You only need a few embryos per staining (but can have the equivalent of 20 µl packed volume if you want). It is a good idea to throw a couple of fly embryos (20-30 or so; rehydrated in PT) into the same tube to act as an internal control. 1. Dissect the embryos out of the female brood pouches. To do this, grab the female around the head or tail with your forceps, and gently nudge the embryos out of the brood pouch with your other forceps. 2. Separate the embryos according to their developmental stage – see below. 3. Dissect individual embryos. For stage 1 embryos, you will need to gently dissect off the egg shell. For stage 2 and older embryos, you will need to separate any embryos that are stuck together. If possible, sonicate these stage 2 and older embryos in PT before beginning the staining protocol to improve antibody access.

Staging: Stage 1: Eggs are spherical. The embryo starts as a small band of cells halfway around the equator of the embryo and then extends into a flattened germband hat is wrapped around the yolk. Once you remove or at least loosen the outer membrane, these embryos need to be dissected only minimally or not at all. If the embryo, which at this point is only a thin sheet, comes loose off the yolk it will float during the antibody staining washes, and you will need to allow more settling time. Stage 2: Embryos appear as very elongated triangles with limbs packed down tightly across the body (antennae may stick out). At this point the embryos have burst out of the egg membranes and will be adhering to one another quite strongly. You usually need to sacrifice the middle one to get the others with all of their limbs, etc. Sonicate these for ~3 seconds. Stage 3: Limbs are now held out to the sides. Clear head and tail are visible. White eyes on eyestalks. Sonicate for ~9 seconds. Stage 4: Brown/red coloring appears in the eyes. For both stage 3 and stage 4 sonicate for ~9 seconds. Staining can be patchy/light because of cuticle.

21 D. 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. 22 E. Parhyale

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 amphipods 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, hold the animals at its posterior half and orient it so that the embryos are facing up at the lens of the dissecting scope. 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 tube, and wash with seawater a few times to get rid of the clove oil. f. To wake mother amphipods up, just 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. pipetteman tip box lined with wet paper towels) on a bench top, or a 26 degree incubator.

Dissecting and fixing Parhyale embryos You will need: Forceps (optional) Plastic Petri dishes (one with Sylgard) Dissecting needles Medicine cups (optional) Fixative (see protocol) PT to rinse fixed embryos 23 Eppendorf tubes Glass Pasteur pipettes and/or plastic transfer pipettes Helpful setup tips: • Dissect embryos in one Petri dish with Sylgard and then transfer them to another dish to fix while you dissect more embryos in your starting dish. • 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 tiny hole in the eggshell with a dissecting needle. Attempt to poke a shallow hole to 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 to 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. 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.

24 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 fixed 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.

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.

25 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.

26 IV.6 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. 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. 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. Fixation and Staining: 1. Fix wing 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. 27 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. Rinse 2-3X in PBS to get rid of detergent. 10. Mount in 80% glycerol. Transfer carcasses/mouth hooks to slide in glycerol and dissect off the desired (wing) 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.

28 B. Butterfly Wing Imaginal Discs

Dissecting wing discs: 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). 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.) Dissect out wing discs by one of the following methods: Method A: • 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. 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: • Use your forceps to tear the skin 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!

Forewing (L) and hindwing (R) discs of late forth/early fifth instar. Note milky, refractant, white trachea around base. (Anterior, top; posterior, bottom.)

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. (Keep in mind the caterpillars you are working with are very inbred, therefore, they may not have all four wing discs.)

2. Place discs in 4% PFA (you will be performing flourescent stains on butterfly discs) for 15 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).

3. Wash in PT until ready to stain. Follow general Drosophila protocol for antibody stains. Fluorescent secondaries generally work better for discs.

29 IV.8 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

30

General Hints and Notes on in situ hybridization experiments

• The fix to use is 1:1 PBS:10% Ultrapure Formaldahyde • 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). a. 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. b. 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 genotypes and get started. • Map out your desired staining protocol ASAP so you can identify any potential problems a. Note that for some genotypes, there are limited amounts of embryos available, so it’s going to be first-come-first-serve. b. Remember that there are also honeybee and beetle embryos available, as well as probes for each (see probe table above). It turns out that Tribolium embryos behave well when it comes to multiplex fluorescent ISHs. In contrast, bee embryos have rarely worked for us fluorescently, but colorimetric in situs are a bit more reliable. • What experiments should you do? 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. 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.

31 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 α DIG Dk α Sh 555 BIO Mo α BIO Dk α Mo 488 DNP Rb α DNP Dk α Rb 647 FITC Rb α DNP Dk α 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.

32 IV.9 Injection in Parhyale

A. Mating and Fertilization Sexually mature male and female Parhyale hawaiensis form mating pairs in which the males grasp and hold the smaller females with their second thoracic appendages (T2, gnathopods) until mating occurs. The pairs remain in premating amplexus until the female molts. At this time the male deposits sperm into the females paired oviducts and then releases her. Before the female’s new cuticle hardens, she sheds her eggs into a ventral brood pouch through two bilaterally symmetric oviducts fertilizing them in the process. The brood pouch itself is composed of several modified, flattened, and interlacing ventral appendage branches (endites termed oosteogites) from the second through the fifth thoracic appendages (T2-T5). At this time single cell embryos are collected by immobilizing the adult female and opening the ventral brood pouch with forceps. For experimental manipulation of embryos, the TAs will collect mature pairs in amplexus in small dishes of seawater the evening prior to lab. The following day, most of the males and females in amplexus have separated. On the day of injection, you should dissect out the embryos from the gravid females and stage them. Note that newly fertilized embryos are very soft and fragile, so should be dissected carefully from the female. Keep your embryos in a clean 35mm Petri dish full of filtered seawater to minimize contaminants. Based on the stage of your embryos, plan your injection schedule accordingly.

B. Injection Needles Our glass needles are 4in-1.0mm with filaments (WPI cat#TW100F-4) and are pulled using a Sutter P-80/PC micropipette puller. We will have pre-pulled the needles for you, and will show you the options for preparing the tip. Loading needles: For this lab, we will backload needles. Your TAs will show you this and can help you if you have problems loading your needles. We will try to have some needles preloaded for you as well.

C. Preparing and Injecting Needles For injecting on a dissecting scope, we usually use sylgard-coated plates that have troughs made in them to inject embryos. The typical injection settings are: Injection pressure is between 25-40psi. Injection time is usually set to 10-20msec These parameters will primarily depend on the bore size of your needle.

D. Timelapse Imaging of Embryos Typically, we place embryos in a 35mm Petri dish with a small hole drilled on the bottom patched with a coverslip (Mattek). The thin glass wall of the coverslip provides greater optical clarity, particularly on inversion-scope timelapse setups. This dish is filled either with filtered seawater or a slurry of 0.02% agarose in filtered seawater and film them in this solution. However, you can also try or devise new techniques. For example, Parhyale embryos will survive “squashing” with a timelapse dish or slide and a coverslip for a limited amount of time (overnight, maybe longer). Your TA’s can help you figure out the best method available for what you want to do.

33 E. Labeling and Lineage Studies A. To visualize germband row formation (ectoderm): • Heat Shock NLS-DsRed transgenic embryos about 12 hours before row formation OR Inject 1/1 or 1/2 cells with DsRed-NLS mRNA • Follow the formation of the germband rows. Can you see the different mitotic waves of division? Can you identify the rows that will express engrailed? B. To visualize muscles: • Collect embryos from the muscle-DsRed line—observe! • Is DsRed expressed in all of the muscles? Double stain with phalloidin and anti-DsRed. C. Lineage studies: • The lineage of the 8-cell stage Parhyale embryo has been previously described (Gerberding et al 2002). See figure in staging section. Repeat some of the lineage studies (using TRITC, or FITC dextran, or DsRed mRNA) and track the movement and fate of different cells via timelapse. • Lineage trace at the 16-cell stage. • Inject Mavg at the 4-cell stage. These cells give rise to the germline and visceral mesoderm. Follow them during gastrulation to visualize the rosette that marks the future anterior pole. • Inject ml or mr with DsRed mRNA, TRITC, or FITC and follow the divisions of the mesoteloblasts. What other tissues do ml or mr contribute to? • Inject ml with FITC and mr with TRITC and follow mesoteloblast development on both sides. How in sync are the two sides? • Combine labeling and staining experiments; antibodies against DsRed and Fluorescein (for FITC-dextran) are available for use.

F. Ablation Studies Given the restricted lineage of the Parhyale embryo, what happens when you kill one blastomere at the 8-cell stage? We know that ablation at the 8-cell stage of one of the mesoderm blastomeres (ml, mr or mav) can be compensated for by the remaining mesoderm. Likewise, after ablation of an ectoblast the remaining ectoderm can compensate for each other. However, ablation of the progeny of these blastomeres at the germband stages does not result in compensation. We use photoablation with FITC-dextran to kill cells. Injecting FITC at a high concentration (25- 50mg/ml) and then irradiating with blue light (ie, GFP filter) will kill a cell or group of cells by releasing free radicals. This method lets you control the timing of ablation.

• Ablate at one of the following times and see if a normal embryo develops: 8-cell stage (as in the original ablation experiments) 4-cell stage late gastrulation (24-28hrs) or early germband (60-72hrs) • Ablate El or Er at the 8-cell stage, and label Ep with NLS-DsRed mRNA, or FITC- or TRITC- Dextran. Does Ep compensate for El or Er at germband stages? • Ablate Mavg and follow how the other cells respond and if the embryo gastrulates properly. Is there a rosette? • Ablate ml and/or mr at different times of development in the muscle-DsRed line. Since ml and mr give rise to most of the muscles, you can use this line to observe if and when ml and/or mr are compensated for.

34 G. Heat Shocking Parhyale 1. Fill an eppendorf tube (if you are using a 37°C water bath) OR a Petri dish 3/4 full with FSW (if you are using a 37°C air incubator) 2. Put this container in the 37°C incubator for about 5 to 15 minutes to allow the water to warm to 37°C. 3. Use a pipet to add your embryos to the 37°C water (these embryos should contain a Hsp70 driven gene, such as PhHsp70:DsRed-NLS) 4. Heat shock at 37°C for 1 hour 5. After 1 hour, remove embryos from 37°C water and place in RT or 25°C water

Notes: For PhHsp70:DsRed-NLS embryos, you should see red fluorescence weakly after 1-2hrs, but strongly after 5hrs if heat shocked at early germband or late germband stages. -You should keep in mind that heat shocking before 24hrs of development does not work. -Once heat shocked, the fluorescence should last for a couple of days (feel free to HS again (maximum once per day*) if you want the fluorescence to last longer). -If Heat shocking the same embryo for multiple days, it may be better to use the air incubator

35 V. General Solutions V.1 Fixatives Formaldehyde Fixes: PEM: 100.0mM PIPES (Disodium salt, Sigma Cat. No. P-3768) 2.0mM EGTA 1.0mM MgSO4 Weigh out the solid PIPES, EGTA, and MgSO4 into a beaker, add the appropriate volume of dH2O, mix for 20 min., adjust the pH to 7.0 with concentrated HCl. The free acid form of PIPES is more difficult to get into solution and the pH will need to be adjusted with NaOH instead of HCl. PEM can be stored for at least one year at 4°C.

Fix 1: 9:1 fix = 9 parts PEM (room temp): 1 part 32% Formaldehyde (PEM-FA) OR 9:1:1 fix* = 9 parts PEM (room temp): 1 part 10xPBS: 1 part Formaldehyde

PEM: 100mM Pipes 150mL = 30mL 0.5M Pipes 2mM EGTA 0.6mL 0.5M EGTA

1mM MgSO4 7.5mL 20mM MgSO4 Adjust pH to 6.95 with NaOH or HCl

Store 4°C Fix 2: 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

V.2 Solutions for antibody protocols 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. 36

PT + 5%NGS: 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).

DAB solution: 1X PBS 0.05% Tween-20 (Sigma Cat. No. P-1379) 3 mg/ml DAB (3,3’-diaminobenzidine; Sigma Cat. No. D-5905) The 10 mg DAB tablets sold by Sigma are very convenient and help minimize the risk of exposure. Note that DAB is a potential carcinogen and should be handled and disposed of in accordance with University regulations. Add one 10-mg DAB tablet to a 50 ml tube containing 33.0 ml PBS and 16.5 µl Tween-20. Rock gently in the dark for about 30 min. Filter through a 0.22 µm filter to remove particulate matter. Store aliquots at –70°C or in a non-defrosting –20°C freezer. Aliquots should be used immediately after thawing.

DAB + Ni solution: Prepare an 8% solution of nickel chloride (NiCl2•6H2O; Fisher Cat. No. N54-500) in dH20. This 8% solution can be stored indefinitely at room temperature. Prepare the DAB+Ni solution by combining 1 ml of the 0.3 mg/ml DAB solution described above with 8 µl of 8% nickel chloride. Mix well and use immediately. It is not advisable to store DAB containing nickel chloride because the nickel will precipitate out of solution (as nickel phosphate) after a few hours.

AP (alkaline phosphatase) buffer: 5.0 mM MgCl2 100.0 mM NaCl 100.0 mM Tris, pH 9.5 0.1% Tween-20 (Sigma Cat. No. P-1379) Prepare just prior to use. The solution will become cloudy after a few hours and then will not work as well for the enzymatic reaction.

BCIP/NBT solution: 1.0 ml A.P. Buffer 4.5 µl NBT (50 mg/ml in 70% DMF) 3.5 µl BCIP (50 mg/ml in 70% DMF) Mix just before use. The NBT and BCIP solutions can be purchased together from Promega (Cat. No. S3771).

37 AEC Buffer:

23.75 ml H2O 400 µl 3M NaAcetate pH 5.2 50 µl 20% Tween-20

AEC Reaction Mix 475 µl AEC Buffer 25 µl AEC in DMF (add AEC/DMF slowly to buffer while mixing). To prepare AEC in DMF, dissolve 20 mg AEC in 2.5 ml of DMF (dimethylformamide), aliquot and store out of light at room temperature.

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%, 70%, and 90% 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 V.3 Solutions for in situ hybridization

PT (1L): 1x PBS 995mL 20% Triton X-100 5mL

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 38

Probe Solution (500ul [1:500]): RNA probe 1ul Hyb Buffer 500ul adjust concentration accordingly

Antibody Solution (3mL): anti-DIG antibody 1ul

39

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 have two pieces of what used to be a long glass pipette. 3. With the larger 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.

40 VII. Available Reagents

VII.1 Antibodies (see separate sheet)

41 VII.2 In situ Probes

Gene DIG BIO DNP FITC Krupple intron X X Knirps intron X X Hunchback intron X X Hunchback 3' X X Antp 3' X X Antp 5' X X X Antp intron X X X X Hairy 5'-intron X X Ubx 3' X X X X Ubx 5' X X Eve X X ftz X X Tailless X X Bxd 3' X Bxd 5' X X sim P1 X X X sim P2 X X CadN 5' X X CadN 3' X X Dfd X Abd-B X X X Abd-A (promoter) X X X Scr (intronic) X Caps X miR-10 X miR-iab-4-3 (S) X miR-iab-4-5 (AS) X D.vir mir-10 X D.vir mir-993 (S) X D.vir mir-993 (AS) X Scr1* X X X X AX1* X X X X AX2* X X X X AC* X X X X AT* X X X X

* Scr1 - SP6 probe detects sense transcripts from the 3' end of the second intron in Scr. - T7 probe is anti-sense

42 AX1 - T7 probe detects sense transcripts from 3' of the second intron of AntpX. - SP6 probe is anti-sense

AX2 - SP6 probe detects sense transcripts from the intron of AntpX. - T7 probe is anti-sense

AC - SP6 probe detects sense transcripts from the putative PRE located between AntpX and the 3' end of Antp. - T7 probe is anti-sense

AT - T7 detects sense transcripts from 3' of the P2 promoter of Antp. - SP6 probe is anti-sense

Note: Sense probes are marked on tube with •

VII.3 Fixed Embryos

D. melanogaster – various ages fixed 15-20 minutes for antibody staining. D. melanogaster w1118 D. virilis T. castaneum A. mellifera

Fixed mutants: (primarily Antp complex mutants)

809 y[1]; Scr[W] Scr[4] p[p]/TM3, Ser[1] 2192 Scr[W] Sb[sbd-2] Ubx[bx-3] Ubx[pbx-1]/TM1 3408 T(3;4)Scr[P], Scr[P]/TM3, Sb[1] 3398 Scr[Wrv3] red[1] e[1]/TM3, Sb[1] 2007 T(2;3)Scr[Wrv1], red[1] Scr[Wrv1] e[1]/TM3, Sb[1] 2189 In(3LR)Scr[9], Scr[9] red[1] e[1]/TM3, Sb[1]

43 VII.4 Drosophila Stocks Reporter Lines his-GFP Histone GFP Drcad-GFP Cadherin GFP Moesin-GFP Moesin GFP (all cell outlines) en-GFP Engrailed neuron MLI-GFP twist-GFP Twist (mesoderm) ubi-GFP.nls; ubi-GFP.nls ubi-DECadherin-GFP DE cadherin Breathless-GFP Tracheal system Kr-lacZ H2Av DGFP histone-GFP Histone-GFP

Crosses hh-GAL4 X GTRACE nub-GAL4 X GTRACE dpp-GAL4 X GTRACE w; FRT82 scrib/TM3 X y, w, ubx-FLP; FRT82B ubi-GFP only females will carry FLP w; FRT82B x y, w, ubx-FLP; FRT82B ubi-GFP only females will carry FLP w; FRT82B wts[x1]/TM6b x y, w, ubx-FLP; FRT82B ubi-GFP only females will carry FLP tub-GAL4 x UAS-apoliner act-GAL4/CyO Act-GFP x UAS-apoliner

GAL4 Lines hh-GAL4/TM6b Hh wg-GAL4 Wg P{w[+mC]=GAL4-elav.L}2/CyO neural Gal4 w; arm-GAL4/TM3 ftz-lacZ ubiquitous Gal4 y[1] w[*]; P{w[+mC]=GAL4-twi.2xPE}2 Twist hh-GAL4, UAS-GFP/TM6b Hh-GFP nub-GAL4, UAS-GFP nubbin (wong disks) P{w[+mC]=UAS-Dcr-2.D}1, w[1118]; P{w[+mW.hs]=GawB}nubbin-AC-62 w[*]; P{w[+mC]=GAL4-btl.S}2, P{w[+mC]=UASp-Act5C.T:GFP}2 tracheal membranes w[*]; P{w[+mC]=RN2-GAL4}E, P{w[+mC]=UAS-tau-lacZ.B}3 eve-GAL4 w[*]; P{w[+mC]=RN2-GAL4}P, P{w[+mC]=UAS-mCD8::GFP.L}LL5 eve-GAL4 w*; wgSp-1/CyO; P{GAL4-dpp.blk1}40C.6/TM6B, Tb1 Dpp-Gal4 w[1118]; P{w[+mC]=GAL4::VP16-nos.UTR}CG6325[MVD1] nanos-GAL4 he-GAL4, UAS-GFP.nls GFP in hemocytes eater-AL4, UAS-2xEYFP GFP in hemocytes

UAS lines UAS-dsRED, UAS-FLP, ubi>stop>GFP GTRACE y[1] v[1]; P{y[+t7.7] v[+t1.8]=TRiP.JF01637}attP2 UAS-N RNAi w[*]; P{w[+mC]=UAS-N.dsRNA.P}14A UAS-N RNAi w[1]; P{w[+mC]=UAS-Ubx.Ia.C}36.2/TM3, Ser[1] UAS-Ubx w[1118]; P{w[+mC]=UAS-Kaede.A}3 Photoconvertible GFP w; UAS-UbxIaC/TM3 ftz-lacZ UAS-Ubx w[*]; P{w[+mC]=UAS-mCD8.ChRFP}3 UAS-membrane Cherry RFP

44 Mutants Box 1 P{ry[+t7.2]=ftz/lacC}4; h[41]/TM3, Sb[1] h ftz nev/o/tau-lacZ ftz P{ry[+t7.2]=ftz/lacC}1, ftz[13]/TM3, Sb[1] ftz y[1]; ftz[11] red[1] e[1]/TM3, Sb[1] ftz th[1] st[1] kni[ri-1] bcd[6] rn[roe-1] p[p]/TM3, Sb[1] bcd kni[ri-1] bcd[15] e[1]/TM3, Sb[1] bcd P{ry[+t7.2]=ftz/lacC}4; hh[21]/TM3, Sb[1] hh ptc[9] cn[1] bw[1] sp[1]/CyO ptc dp[ov1] b[1] BicD[1]/CyO BicD dp[ou1], b[1], Bic-D[1}/CyO BicD Dl[B2] e[1]/TM6C, Sb[1] Dl w[ch2] N[264-39]/FM4, B[+] N N[55e11] P{ry[+t7.2]=neoFRT}19A/FM7c N ru[1] h[1] th[1] st[1] cu[1] sr[1] Dl[9P] e[s] ca[1]/TM3, Sb[1] Dl w[*]; lola[ORE119]/CyO lola y[1] w[*]; robo[1]/CyO, P{w[+mW.hs]=ase-lacZF:2.0}PK2 robo w[*]; robo[2]/SM6b robo Slit/CyO (enII) wg-Bgal Slit y[1] w[67c23]; P{w[+mC] y[+mDint2]=EPgy2}comm[EY10154]/TM3, Sb[1] commissureless Ser[1] Box 2 P{ry[+t7.2]=ftz/lacC}1, opa[8]/TM3, Sb[1] opa P{ry[+t7.2]=ftz/lacC}4; opa[8]/TM3, Sb[1] opa P{ry[+t7.2]=ftz/lacC}1, opa[5]/TM3, Sb[1] opa odd[5]/CyO; P{ry[+t7.2]=ftz/lacC}1 odd prd[9] cn[1] bw[1] sp[1]/CyO prd w[*]; prd[9]/CyO; P{ry[+t7.2]=ftz/lacC}1 prd prd[II], B[42], en, bw, sp/CyO prd en[59]/CyO; P{ry[+t7.2]=ftz/lacC}1 en w[*]; Df(2R)en-B, b[1] pr[1]/CyO en eve[1]/CyO; P{ry[+t7.2]=ftz/lacC}1 eve hs eve 19B/CyO hs-eve hs eve 19B/hs eve 19B hs-eve eve R13 bpr/CyO eve cn[1] bw[1] sp[1] Kr[2]/SM1 Kr Kr[2]/CyO; P{ry[+t7.2]=ftz/lacC}1 Kr y[1] w[*]; wg[l-8] cn[1] bw[1] sp[1]/SM6b, P{ry[+t7.2]=eve-lacZ8.0}SB1 wg wg[1] cn[1] wg wg[l-8]/CyO; P{ry[+t7.2]=ftz/lacC}1 wg wg[ts]/CyO Belav wg temperature sensitive allele Box 3 Ubx[bx-3] Ubx[bxd-1] Ubx[pbx-1]/T(2;3)ap[Xa], ap[Xa] (2, 7) 4 winged flies gsb[525]/CyO ftz-lacZ weak gsb allele (CNS phenotype) w2a w-; GA881/CyO Belav weak gsb allele (CNS phenotype) w; FRT82B crb[269]/TM6b w; FRT82B crb[163]/TM3 GFP w; FRT82B wts[2H270]/TM6b w; FRT82B scrib[2], e/TM3

45 VIII. DEVELOPMENTAL BIOLOGY & ANIMAL STAGING

VIII.1 Parhyale (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 discreet 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.

46 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.

47 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

48 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.

49

V

50 VIII.2 Drosophila

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

51 Here is a schematic diagram that you can use to identify fly embryo developmental stages:

52

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)

53

54 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.

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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.

56 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 characterised, 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.

57 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

58 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.

59 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.

60 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 satge 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.

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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.

62 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 are 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.

63 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 GFP. 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).

64 VIII.3 Spiders

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.

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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.

66 VIII.4 Grasshoppers

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)

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