/ Lab 2009 June 22-27

Lectures: Nipam H. Patel [email protected] (Mon & Fri) Steve Small [email protected] (Tue) Matt Ronshaugen [email protected] (Wed) Andrea Brand [email protected] (Thur)

Labs: Andrea Brand [email protected] Cassandra Extavour [email protected] Pan-Pan Jiang [email protected] Henrique Marques-Souza [email protected] Nipam H. Patel [email protected] Matthew Ronshaugen [email protected]

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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.

Possible “Projects”

1) Segmentation Look at the mutant cuticle phenotypes of Drosophila segmentation mutants. Examine the expression patterns of segmentation genes in Drosophila (protein and mRNA), and compare them to the patterns of orthologous genes in orther (for example, examine engrailed and even- skipped expression in multiple species) Carry out live imaging of GFP lines for segmentation genes in Drosophila

2) Hox genes Look at the effect of Hox gene misexpression in Drosophila using the Gal4 UAS system Examine the expression patterns of Hox genes in Drosophila (protein and mRNA), and compare to Hox gene expression in other arthropods. Look at the expression of non-coding mRNAs from the Hox complex

3) Neurogenesis Look at the process of neurogenesis in Drosophila using various antibodies Look at mutants that effect fly neurogenesis Compare neurogenesis in multiple arthropod species. Examine neurogenesis in the fly eye disk

4) Axonogenesis Look at patterns of axonogenesis in Drosophila Examine mutations causing defects in axonogenesis Watch living neurons pathfind to their targets Examine the pattern of motoneuron connectivity with muscles in larvae, and axonogenesis in the eye disk. Compare axonogenesis patterns in multiple species

5) Dorso-ventral patterning Look at Drosophila mutations that effect D/V patterning Look at the expression of genes involved D/V patterning in Drosophila and compare to expression patterns in other arthropods. Look at the role of single-minded in Parhyale D/V patterning

Page 2 of 80 6) Organogenesis Watch organogenesis in living Drosophila embryos using GFP expression Look at the expression pattern of genes involved in patterning organs such as somatic miscles and the heart and compare this to the patterns you see in other arthropods.

7) Appendage/Disk development Look at the development of Drosophila imaginal disks. Compare patterns of gene expression between Drosophila imaginal disks, the imaginal wing disks of butterflies, and the appendages of arthropods that develop directly without imaginal disks. Use Gal4 UAS system to lineage trace cells from the embryo into the disks

8) Germline/ovaries Examine the specifications, development, and migration of the germline in various arthropods. Examine ovary development in Drosophila and compare to ovary development in Triboloium, Oncopletus, and Schistocerca.

Page 3 of 80 II. SCHEDULE

Monday, June 22 Lecture: Nipam Patel: Intro to Arthropod Development Afternoon and Evening Labs: Arthropods/ 1 and 2 Goals: Start antibody stains on Drosophila embryos Be able to stage and navigate the Drosophila embryo Plan general experiments for the rest of the week Start fly crosses and in situs

Tuesday, June 23 Lecture: Steve Small Afternoon and Evening Labs: Arthropods/Flies 3 and 4 Goals: Continue antibody and in situ stains on Drosophila embryos Learn to dissect many other arthropods (and start staining!)

Wednesday, June 24 Lecture: Matt Ronsgaugen Afternoon Lab: Arthropod/Flies 5 plus Plankton Tow/Outdoor Collecting (snorkeling!) Evening Lab: Arthropod/Flies 6 Goals: Find embryos to examine in Plankton Tow or from your own collecting Continue other experiments

Thursday, June 25 Lecture: Andrea Brand Afternoon and Evening Labs: Arthropods/Flies 7 and 8 Goals: Continue working on dissections, injections, stainings

Friday, June 26 Lecture: Nipam Pateln Afternoon and Evening Labs: Arthropods/Flies 9 and 10 Goals: Continue working on dissections, injections, stainings

Saturday, June 27 Lecture: Richard Harland Afternoon Lab: Arthropods/Flies 11 Goals: Finish Arthropod/Fly projects and get ready for Show and Tell Evening: Show and Tell!!!

Page 4 of 80 III. Experimental Details Section 1. Antibody Staining a. Drosophila 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. Each group should complete all seven of the following antibody stains on Drosophila. You will be using a “Rapid” version of the standard antibody.

Your fly embryos are a mix of embryos ranging from 0-17 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. Your aliquot should last you throughout this and future experiments during the Arthropod module. Additionally, too much tissue will give a decrease of signal!

React Antibody Staining Pattern µl:300 1:3000 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 Goat anti-mouse regional expression pattern Ni HRP (115-035-003) DAB+ 3) BP102 Stains all CNS axons (unknown antigen) 1.5 Ni 4) BP102 See above 1.5 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 7) Goat anti-HRP-FITC 2.0 NO SECONDARY flour

Page 5 of 80 While this protocol produces antibody stains in one day, it only works well on very robust antibodies. The “usual” protocol can be found further down in this manual.

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 1:250. 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.

Page 6 of 80 Into the wild

Use the protocol and expertise 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 ) Schistocerca americana (grasshopper) Apis mellifers (honeybee) Oncopeltus fasciatus (milkweed bug)

Crustaceans: Parhyale hawaiensis (beach hoppers—amphipod crustaceans) Gammarus sp. (beach hoppers—amphipod crustaceans) Orchestia species Mysidium columbiae (opossum shrimp) Triops longicaudatus (tadpole shrimp) Artemia salina (brine shrimp or “sea monkeys”) Marmokrebs. (marble crayfish)

Chelicerata: Parasteatoda tepidariorum (common house spider)

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Molecular markers of embryonic development

c.1. 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 (and 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) on Parhayle

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

c.2 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, Oncopeluts, Grasshoppers (15 – 30%), Parhyale, Mysids, stage 1, 2, and 3 (sonicate stage 2 and 3), Triops and 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 Triops and 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)

Page 8 of 80 c.3. 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), Triops (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 c.4. 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-Alk Phos on Artemia, Triops, and stage 3 Mysids (all sonicated) (develop w/BCIP/NBT) 11) Goat anti-HRP-Alk Phos on Drosophila (develop w/BCIP/NBT) 12) Goat anti-HRP-Alk Phos on Grasshopper (develop w/BCIP/NBT) 13) Goat anti-HRP-Alk Phos on Parhyale and Spiders (develop w/BCIP/NBT)

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 **Make certain to carry out “no primary” controls on the anti-HRP Alk Phos stains for each species!** c.5. Neurogenesis and Axonigenesis Fluorescently! Triple label the fly CNS with antibodies made in different animal species (mouse, rabbit, rat). 1) Mouse DP312 (anti-Pax3/7+) GαM 555 OR 4D9 (anti-engrailed) GαM 555 2) Rabbit 10900 (anti-eve) GαRb 647 3) Rat 7E8 (nuclei of all neurons) GαR 488 OR anti-HRP FITC (all neurons) (no secondary needed) Page 9 of 80

a. Additional genes

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

b. Specific tissue stains e.1. WING IMAGINAL DISK 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 disks (3rd instar) 2) 4F11 (anti-Engrailed) on buckeye wing disks (4th and 5th instars) 3) FP6.87 (anti-Ubx) on Drosophila wing disks (3rd instar) 4) FP6.87 (anti-Ubx) on buckeye wing disks (4th and 5th instars)

Double label: 5) 4F11 (mouse 488) + anti-Spalt (rabbit 546) on Drosophila wing disks (3rd instar) 6) 4F11 (mouse 488) + anti-Spalt (rabbit 546) on buckeye wing disks (5th instars)

Notes: You can stain Drosophila and butterfly disks in the same tube Put fly embryos in as controls (will stain for Engrailed and Spalt) Use fluorescent secondaries on disks (while Drosophila disks will stain fine histochemically, butterfly wing disk staining works best flourescently). Only late 5th instars and older butterfrly larvae will have eyespot staining. For best eyespot stains, use disk from 12hrs post pupariation. e.2. OVARY STAINING vasa to highlight the germline, phosphotyrosine to see connections between nurse cells. Hunchback to see the position of the oocyte nucleus in grasshoppers.

Page 10 of 80 SUGGESTED Drosophila EXPERIMENTS

The following are suggestions only: please feel free to invent your own experiments or assays as well, and ask the TAs for help designing these if you need it.

Please don’t try to do everything! Instead, choose those aspects of development, or techniques, most interesting to you.

If you plan to do experiments involving in situs, antibody stainings, or crosses (GAL4/UAS), we suggest that you make a chronological plan before starting, so that you know when plan your crosses, incubations and washes, and to leave you enough time to analyze them on subsequent days.

Ask TAs for help if unsure.

Observations of embryogenesis • Antibody Staining: You can antibody stain pre-fixed mixed stage wild type embryos of Apis mellifera (take your chances with cross-reacting antibodies), Drosophila melanogaster, or Tribolium castaneum to examine particular cell types or signaling molecules • Live Imaging of Wild Type 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 (Cassandra will demonstrate mounting for long term (>30 minutes or so) time lapse). o Wild type o GFP lines (e.g. moesin:GFP, histone:GFP…) o GAL4 lines crossed with UAS:GFP (e.g. engrailed:GAL4, armadillo:GAL4) • 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! Here are some of the mutant stocks we have available (for a full list see p. 57): o (1) bicoid (bcd) mutant: we have a loss of function alleles of this maternal gene o (1, 3) engrailed (en) mutant: a segment polarity gene o (1, 3) even-skipped (eve) mutant: a pair rule gene o (1) folded gastrulation (fog) mutant o (1) fushi tarazu (ftz) mutant: a pair rule gene o (1) Kruppel (Kr) mutant: a gap gene o (1, 8) nanos (nos) mutant: a posterior maternal determinant o (1) wingless (wg) mutant: a segment polarity gene o (1) knirps (kni) mutant: a gap gene

Combinatorial Fluorescent Antibody Staining • Assay localization of several proteins simultaneously • You are only limited by the 1° antibodies and fluorescent 2° antibodies available (check with TAs and consult the list in the back)

Page 11 of 80 (1, 2) 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 and Tribolium. • Analyze the denticle belt patterns for embryonic patterning mutants: e.g. Kr, eve, ftz, wg, en, bcd, and nos • You can also collect embryos from these crosses and stain them with various antibodies.

GAL drivers in development, in live or fixed • Express GFP or RFP in specific tissues by crossing GAL4 and UAS::GFP lines • Express Ubx ectopically in specific tissues by crossing GAL4 and UAS lines • Observe live during development (look at GFP), or fix and stain with interesting antibodies to observe GFP plus expression of another gene o Look at ventral furrow formation using DEcadherin-GFP o Observe early nuclear division cycles using GFP-nls

(7) Larval structures/Imaginal disc development • Stain imaginal discs using antibodies against Wingless, Engrailed, Armadillo, etc., and observe changes in gene expression patterns through development. • Squash larval brains and/or salivary glands to do cytological analysis: o Polytene chromosomes: identify chromosomal regions and balancer chromosomes by their banding pattern

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

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Parhyale RNAi and Live-Imaging Experiments

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 to lineage tracing and ablation experiments. Additionally, we have brought two transgenic lines that will facilitate your observations of Parhyale development. You can also try knockdown of single-minded via siRNA injections.

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 Anti-DsRed (Rb) Nuclear-localized red fluorescent Use b/w 500ng/ul to Use at final conc. of protein- will label all progeny of 1ug/ul 1:1000 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 — Red fluorescent dye- able to visualize (Tetramethylrhodamine Use at 25mg/ml-50mg/ml immediately isothiocyanate–dextran) for lineage tracing

HS-NLS-DsRed Parhyale Heat shock embryos: Anti-DsRed (Rb) Nuclear-localized red protein- after (transgenic line produced by 1 hour at 37°C/day Use at final conc. of heat shock, will appear in all cells M. Modrell, HS promoter from 1:1000 T. Pavlapoulos) Muscle-DsRed Parhyale A muscle promoter drives Anti-DsRed (Rb) Red protein in muscles (transgenic line produced by DsRed in this line. Use at final conc. of R. Parchem, muscle promoter Sit back and watch! 1:1000 from T. Pavlapoulos)

CAN ALSO TRY LASER “HEAT SHOCK” TO TRACE CELLS

Page 13 of 80 IV. Protocols IV.1. General Antibody Staining

Note - If embryos were fixed today and not dehydrated in MeOH, skip step 1 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 table in back of manual).

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.

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 conjugated secondary antibodies, proceed to the histochemical development reactions on the next two pages.

Page 14 of 80 Histochemical development reactions: Unless specified, reagents are on BACK BENCH or in your GROUP BOX 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 1.0 ml aliquots have already been made and are in the -20°C freezer. 1.0 ml is enough to do 3 reactions. Thaw the DAB. Once completely thawed, add 8 µl of 8% nickel chloride to 1.0 ml of DAB. Mix by inverting the tube several times. Treat all DAB waste with care. Liquid waste should go into 50 ml Falcon tubes clearly labeled as “liquid DAB waste”. All solid waste (used tips and eppendorf tubes) should go into a zip-locked bag labeled “solid DAB waste”.

2. After the last PT wash, drain the tubes down to about 1 mm above the embryos. Add 300 µl of DAB+Ni. 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 3% hydrogen peroxide with 90 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

1. Follow the same protocol as above, but leave out the addition of nickel chloride C. Red HRP reaction

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

Page 15 of 80 Purple Alkaline Phosphatase 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 Alkaline Phosphatase 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.

Page 16 of 80 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 1:250. Mix and incubate in the secondary antibody for 30 min at room temperature.

13. Wash 3X 1 min with PT.

14. Wash 3X 10 min with PT.

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

16. Wash with PT.

17. 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.

Page 17 of 80 IV.6 Injection in Parhyale 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 and other crap. Based on the stage of your embryos, plan your injection schedule accordingly. 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. PREPARING AND INJECTING EMBRYOS For injecting on a dissecting scope, we usually use sylgard coated plates that have troughs made in them to inject embryos. However, the scratched dish technique that you learned previously will also work. 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. Timelapsing 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 sea water 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.

Page 18 of 80 IV.7. Parhyale Labeling and Lineage Studies 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? 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. 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 tissue 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. IV.8. 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 methods let you control the timing of ablation. - Ablate at one of the following times and see if a normal embryo develops: 8-cell stage (via 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.

Page 19 of 80 V. ADDITIONAL PROTOCOLS

V.1. FIXATIVES Formaldehyde Fixes: Fix 1: 9:1 fix = 9 parts PEM (room temp): 1 part Formaldehyde (PEM-FA) 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 37% formaldehyde 9:1 seawater fix* = 9 parts filtered seawater: 1 part 37% formaldehyde

Formaldehye: 37% Fisher cat # F79-500 *saltier fixes are for Parhyale

Paraformaldehyde Fix (PFA) (use for embryos to be fluorescently stained): . Solution A = 2.26% NaH2PO4 H2O Solution B = 2.52% NaOH

Solution C = 1.0% CaCl2

50mL= 1) Dissolve 2g of PFA into 8.5mL Soln B Stir until clear; Do not heat! 2) Gradually add 41.5mL of Soln A 3) Add 0.6g of glucose 4) Gradually add 0.25mL of Soln C 5) PH to 7.3 with pH paper 6) Filter sterilize and store at 4°C (good 2-3 weeks)

Notes: 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 the fixed embryos from sticking to the pipette. You can introduce detergent in the last 2-5 minutes of fixation by adding 1/5 volume of PT to your fixative.

V.2. DEHYDRATION AND STORAGE

Embryos can be dehydrated through a MeOH/PBS series and stored in 100% MeOH at -20°C. Page 20 of 80 V.3. GENERAL 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.

PT+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).

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.

PEM-FA 9.0 ml PEM 1.0 ml 37% formaldehyde (Fisher Cat. No. F79-500; this can be stored at room temperature for at least one year) This PEM-FA solution should be made just before use. For immunohistochemical procedures, it is unnecessary to start with solid paraformaldehyde. The 37% formaldehyde solution is stabilized with 10-15% methanol, however, so when using the protocols to detect antigens sensitive to methanol, the fixative should be prepared from solid paraformaldehyde.

DAB solution 1X PBS 0.05% Tween-20 (Sigma Cat. No. P-1379) 0.3 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.

A.P. (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).

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 Page 21 of 80 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

Page 22 of 80 V.4. LABELING WITH MULTIPLE PRIMARY ANTIBODIES

Antibodies made in different animals (fluorescent and histochemical) If all of the antibodies are made in different animals (for example: mouse, rabbit and/or rat), then you can do all of your stains at the same time. Additionally, 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 substrates, first add your primaries at the same time, then add the first secondary and perform that reaction; followed by adding the second secondary and performing that reaction.

1° step: add the appropriate amount of each primary antibody to 300µl PT+5%NGS

2° step: add the appropriate amount of each secondary antibody to 300µl PT+5%NGS

color reaction step (for histochemical stains): perform the color reactions sequentially. Make sure to let you reactions go to completion—at least 10 minutes. Wash well with PT b/w reactions!

Antibodies made in the same animal (histochemical) If the 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, do the first reaction rapidly (see protocol section IV.2), if your first antibody is particularly robust. This rapid protocol can then be followed by a regular length protocol(s). If all of your antibodies are not-so-robust, then follow a regular length protocol with another regular length protocol(s). Examples of robust antibodies are BP102, 22C10, and DP312. Others are noted in the suggested experiments, but feel free to ask about additional antibodies.

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”.

Page 23 of 80 V.5. PROTOCOLS FOR FIXING AND STAINING ARTHROPODS

V.5.1 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: 37% 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.

Page 24 of 80 V.5.2. ARTEMIA AND TRIOPS These crustaceans require special permeabilization techniques for optimal staining.

Protocol: A. FIXATION 1. Fix animals in 9:1:1 FSW OR PEM:10X PBS:37% 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 (section C) or dehydrate into methanol gradually and store in 100% methanol at –20C.

B. REHYDRATION 2. 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. 3. Rinse the animals in 100% PT several times (5 or 6 changes).

C. PERMEABILIZATION AND BLOCKING 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 - a few (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. 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 (50 or so; rehydrated in PT) into the same tube to act as an internal control. D. ANTIBODY INCUBATIONS 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. F. 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.

Page 25 of 80 V.5.3. MYSIDS

These embryos have already been fixed for you (fixed while still in the female brood pouch). They had been stored in methanol, and we have already rehydrated them in PT for you (3x10min PT at room temp).

Protocol:

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.

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.

Mysid 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.

Page 26 of 80 V.5.4. 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 V. figure b, white arrowhead). 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!**

Putting amphipods to sleep: • Add 10uL of clove bud oil to 50mL of filtered seawater in a falcon tube. Shake vigorously. • Collect gravid Parhyale in a Petri dish or a medicine cup. • Remove as much water as possible. • Add your clove oil / seawater mixture (cover the Parhyale). • Wait for them to completely stop moving – 5 to 10 minutes should do the trick. • Caution: Leaving amphipod in the clove oil too long (hours) will kill them.

Embryo extraction: • 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). • 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. • 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. • ** 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!** • Transfer embryos to a new tube, and wash with seawater a few times to get rid of the clove oil. • 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! • Put the adult amphipods back into their tank. Note: Do not place sleeping amphipods back into their tank – they will be eaten by those that are awake.

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.

Page 27 of 80 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 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 love to stick to glass pipettes and even the sides of the wells in the glass dishes. One easy way to prevent sticking is to use some yolk from the first embryo (or group of embryos) you dissect to swish around the bottom of the dish and to suck up into the pipette prior to pipetting the actual embryos. It also helps to only suck embryos up into the narrow neck of the pipette – avoid the wider part where embryos tend to stick more frequently. Protocol: 1. Place a few embryos (start with 2-3 and increase with experience) in the well containing fixative. 2. Holding each embryo in place with your forceps (forceps are optional, you can also position your embryo with one of your dissecting needles), poke a tiny hole in the eggshell with a dissecting needle. Be sure to poke a shallow hole to avoid destroying the embryonic tissues, and try to poke a hole in the yolk away from most of the embryo if possible (see figure next page). This becomes 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 minute – this assists in the dissection but be careful not to wait too long because the embryo is fixing to the membrane(s) as well. 4. Carefully peel away the outer membrane starting at the hole you made with your dissecting needles. Sometimes it helps to make a slight tear in the membrane at the poked hole because it will produce a flap or loose end of membrane you can hold with one needle or forcep. 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 outer membrane! If not, repeat steps 4 & 5 for inner membrane. 7. Try to get the embryo mostly out, and relatively 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 for. Try to remove the membrane within 5 minutes and then allow the embryos to fix for 15 more minutes (total time in fix should be around 15-20 minutes). 8. Once fixed, pipette embryos into a dish/tube containing PT to rinse embryos before beginning the staining protocol. 9. 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

Page 28 of 80 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.

Page 29 of 80 Helpful hints for dissecting different age Parhyale embryos: 0-18hrs: These embryos are very yolky and 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, for antibody staining 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 another fixative (9PEM:1PBS(10x):1Formaldehyde) once the membrane has been totally removed.

1-2 day: These embryos are relatively easy to dissect because they are surrounded by a single membrane and since 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-3day: This is another tricky stage because the embryo is not easy to identify among the yolk has developed two outer membranes – one of which is easy to remove (outermost) and another membrane that is very difficult. 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 less 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 to the embryo. If you cannot remove all the membrane, remove all the yolk on the opposite side – this exposure will be enough for antibody staining to work (probably not in situs though).

4+day: Once the embryo has grown appendages, the trick is to remove the membrane without dismembering. It is easier to dissect many older, leggy embryos if you poke a hole behind their head and antennae – you are less likely to split the embryo through it's abdomen and have more room to tear off some membrane. After poking a hole through the membrane, it may actually be helpful to let them sit for a minute or two before touching them again – limbs may remain more intact if they are fixed for a while before you dissect the membrane away from the embryo.

Page 30 of 80 V.5.5. GRASSHOPPERS

1. Fill a Sylgard coated dish with 1XPBS (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. (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 lots of background, use the secondary at 1:600.

Page 31 of 80 V.5.6. BUTTERFLY WING IMAGINAL DISKS

1. 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). 2. Fill a Sylgard coated dish with 1X PBS. Place one larva in dish. Orient the larva so that dorsal is up (towards you) and ventral is down (towards the Sylgard). 3. 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.) 4. Dissect out wing disks: Method A: 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! b. Carefully pull back the skin on one side; pin down if desired. The imaginal wing disks 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.) c. Locate wing disks: They are a bit posterior and ventral to the lateral bristle. The disks 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 disk. Method B: Use your forceps to tear the skin behind the lateral bristle of the 2nd and 3rd thoracic segments. The imaginal disks will either “pop” out or will be sitting in the nearby tissue. Be careful not to harm the disks when you tear the skin!

Forewing (L) and hindwing (R) disks of late forth/early fifth instar. Note milky, refractant, white trachea around base. (Anterior, top; posterior, bottom.) 5. Dissect out wing disks: carefully tear away the tissue at the base of each disk. Place in 1X PBS until all four wing disks are found. 6. Fix disks: place disks in 4% PFA (you will be performing flourescent stains on butterfly disks) for 15 minutes. While fixing, dissect off remaining trachea and/or tissue from the disks. You can also try to remove the peripodial membrane, although antibody staining works with membranes on (in situs do not). 7. Wash in PT until ready to stain. 8. Follow general Drosophila protocol for antibody stains. Fluorescent secondaries generally work better for disks.

Q: I can’t find one (two, three, four) wing disks in my caterpillar. Where are they? Page 32 of 80 A: The caterpillars you are working with are very inbred, therefore, they may not have all four wing disks.

Page 33 of 80 V.5.7. DROSOPHILA WING IMAGINAL DISKS

1. Fill a Sylgard coated dish with 1X PBS. Place one 3rd instar larva in dish.

2. Dissect out wing disks:

Method A: 1) Cut embryo in half (cut across “waist”) with dissecting scissors 2) Grasp mouth hooks with 1st pair forceps 3) Invert body by pulling on mouth hooks. Use 2nd pair forceps to help. 4) Wing disks will be in a bunch with other disks. Wing disks are largest and have a pocket.

Method B: 1) Grasp mouth hooks with 1st pair forceps 2) Grasp middle of body with 2nd pair forceps 3) Pull mouth hooks (and connected imaginal disks) out of body 4) Wing disks will be in a bunch with other disks. Wing disk is largest and has a pocket.

3. Fix disks: place disks in 4% PFA (if performing a flourescent stain) for 15 minutes. While fixing, remove any remaining extraneous tissue. You may want to keep the wing disk attached to the other disks and/or mouth area in order to have a larger object to keep tract of during antibody staining.

4. Wash in PT until ready to stain

5. Follow general Drosophila protocol for antibody stains.

Page 34 of 80 V.6. ADDITIONAL PROTOCOLS FOR PARHYALE (We encourage you to try these on other arthropods as well!)

V.6.1 MAKING DISSECTION TOOLS

To make a Blunt probe: • Heat a long glass pipette with a Bunsen burner and pull so that the long skinny part stretches and breaks. • Now there are have two pieces of what used to be a long glass pipette. • 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.

To make Tungsten needles for dissecting: (station in back of the main lab)

Electro-chemically sharpened: • Use Tungsten wire 0.005” diameter (Ted , Inc., the Electron Microscopy Supply Center; Redding, CA. 1 • 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. • Set up a beaker with 1 N NaOH. WEAR SAFETY GOGGLES TO PREVENT SPLASHING OF NaOH INTO YOUR EYES! • 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!!! (and you will short out the transformer) • Plug clamps into the Variable Auto Transformer. Set input = 120 V, 50/60 Hz; Output = 0- 120/140 V, 10 A, 1.4 KVA. • Put switch on 120 V. Set dial to at least “2”, but no higher than 6. • 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.

V.6.2 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, soniate 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

Page 35 of 80 V.6.3. HEAT SHOCKING PARHYALE

Protocol: 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 germcap 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 HSing the same embryo for multiple days, it may be better to use the air incubator

Page 36 of 80

V. ARTHROPOD STAGING

V.1. 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

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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.

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.

Page 38 of 80 V.2. PARHYALE

Modified from Browne et al (2005) Genesis 42:124-49 and Gerberding et al, (2002) Development 129:5789-5801.

SUMMARY 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.

Page 39 of 80 Quick reference guide to Parhyale development

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.

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.

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.

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S9-17 Germband formation and elongation (Left panel: live image and matching DAPI (nuclear) images, Right panel: ventral view of embryos stained with Engrailed Brightfield and DAPI images shown) 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 progression of segmentation in the A/P axis.

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S 18-30 Appendage formation, organogenesis and neurogenesis to hatching (Left panel: live image and matching DAPI (nuclear) images, Right panel: ventral view of animals stained with Engrailed, Brightfield and DAPI images shown) 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)

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VII.3. GRASSHOPPERS

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Grasshopper Pax3/7 and Engrailed expression

Page 45 of 80 INTRODUCTION TO DROSOPHILA DEVELOPMENT

SEM of a gastrulating D. melanogaster embryo. Anterior is to the left. 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.

Embryos can be dechorionated, mounted in Halocarbon (or Voltalef) oil, and observed live under oil for a few hours. For longer live imaging, use a bio-foil mounting setup (demonstrated by Cassandra). Transgenic lines can also be used to observe the development and morphogenesis of the embryo or particular cell types. Crossing tissue specific GAL4 lines (e.g. en:GAL4) with UAS:GFP lines is another way to observe the development of specific cell types in live or fixed embryos. Some GFP lines will retain GFP fluorescence after standard fixation protocols, but you can also use an anti-GFP antibody to identify cells expressing GFP in fixed tissue.

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Stages of Embryonic 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

Page 47 of 80 Here is a schematic diagram that you can use to identify fly embryo developmental stages:

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 Page 48 of 80 check out the Drosophila Atlas of Development by Volker Hartenstein, available at the “Interactive Fly” website: see URL p. 45)

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Page 51 of 80 Larval Development, Imaginal Discs, 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.

(A) Fate map of imaginal disc primordia in a blastoderm stage embryo. (B) Position of the imaginal disc primordia at the end of embryogenesis. Anterior is to the left.

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 other genes, play important roles in specifying imaginal disc identity, so 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. You can dissect these discs and stain them with various antibodies and cell biology markers to observe the patterning processes, mitotic proliferation, and changing morphologies.

Top: Position of the imaginal disc in an L3 larva. Bottom: enlarged view of the imaginal discs and gonads of both sexes of an L3 larva. Anterior is to the right.

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 Page 52 of 80 (pupal case) is formed, and the process of metamorphosis begins. Most of the larval tissue is destroyed by histolysis, and 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.

Page 53 of 80 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.

Page 54 of 80 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.

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. For this reason, for best staining, fixation of ovaries is often done in a mix of 1:1 heptane and aldehyde fixative (formaldehyde, paraformaldehyde, or glutaraldehyde). In this lab, our standard fixative will be 3.7% formaldehyde in 1x PBS (without the heptane).

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A pair of D. melanogaster ovaries from an old adult female, still joined at the common oviduct, filled with mature eggs. Anterior is at the top.

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 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.

Page 56 of 80 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. You can stain the fusome and the presumptive oocyte with specific antibodies in this lab to visualize the process of oocyte specification. 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.

Germarium and early oocyte stained with phalloidin and a ring canal-specific protein. Anterior is to the left.

Page 57 of 80

The Male Reproductive System

The testes of adult males also have a stem cell region called the hub at the anterior, but the posterior regions of the testes are organised 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.

Page 58 of 80 Multiplex 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 best to 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 (p.52). • For many genes we have probes for flies, and bees. Why not see how the expression patterns have evolved, and look at your gene of interest in different orders? Evo-Devo fun!

Page 59 of 80 SUGESTED EXPERIMENTS: SOME SPECIFIC D r o s op h il a I N S I T U A SSAY S

1. (1) 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. How about a multiplex in situ to assay what happens to gastrulation in these embryos? 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. (6) 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. (2) 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. (3, 6) Observe the border of the mesoderm and neurectoderm by visualizing rho, sim and snail (or twist). Also you can examine what happens when some of the early mesodermsal DV transcription factors expressed in an AP gradient. 6. (3) Observe the differential expression of the related but differentially expressed genes SoxN and Dichaete in the nervous system1. 7. Compare the expression pattern of midline patterning genes (e.g. sim) between Drosophila and Apis embryos2.

1 Conserved genomic organisation of Group B Sox genes in . McKimmie, Woerfel, Russell. BMC Genetics (2005) 6(1):26 2 Zinzen, Cande, Ronshaugen, Papatschenko, Levine. Evolution of the ventral midline in insect embryos. Developmental Cell (2006) 11(6):895 Page 60 of 80

SUG G ESTED EXPERIMENTS: D r o so p hi l a MULTIPLEX I N S IT U HYBRIDISATION

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.

Page 61 of 80 General Hints and Notes on Drosophila in situ hybridization experiments

- 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).

o 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.

o unless otherwise indicated, washes are not all that time-sensitive, especially when they are pure alcohol or PBT 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

o note that for some genotypes, there are limited amounts of embryos available, so it’s going to be first-come-first-serve.

o 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?…

o 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.

o Do (or get one of your friends to do it and show you) a WT (i.e. yw) control in parallel with mutant genotypes.

- you may want to compare protein and RNA expression patterns. Does the protein, e.g. Twist, exactly overlap the area where the gene is transcribed?

- you could compare genes that are supposedly co-expressed in the same domain (e.g. vNE), or you could compare transcription factor distributions (i.e. Dorsal, Twist and Snail), and quantify relative distributions along the DV axis on the confocal.

Page 62 of 80 - rhomboid is expressed in a lateral stripe in the ventral neurogenic ectoderm at first, but becomes stripy just before cellularization completes. You could combine rho and eve stains to determine whether this AP modulation conforms to even or odd parasegments.

- Suggested probes and antibodies (pick a combo): sim, rho, SoxN, slit, otd, brk, vnd, ind probes with Elav antibody - What happens when you mess with the Dorsal gradient? For example:

o -Tollrm9/rm10 (hypomorphic Toll) for neurogenic ectodermal genes (vnd, ind, rho, brk) and a mesodermal marker (Twist, Snail)

- 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.

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AVAILABLE PROBES FOR DROSOPHILA I N S IT U HYBRIDISATION

These are shared reagents in labeled boxes at -20ºC Boveri freezer. Ask TAs for help as we have limited quantities. Please make sure you have your entire staining scheme mapped out before you add probes.

PROBE SYMBOL DIG DNP BIO NOTES RATING Mesoderm heartless htl x x x mesoderm  twist twi x mesoderm  cactus cact x dorsal/ventral axis specification  tinman tin x x heart marker, pan mesodermal early  Myocyte enhancing factor 2 Mef2 x x late mesodermal marker only  bagpipe bap x x x visceral mesoderm  biniou bin x heart marker  snail sna x x probably intronic  Neuro-Ectoderm tartan trn x x DV AP criss-cross pattern  ventral nervous system defective vnd x ventral neuroblasts  ventral nervous system defective intronic vnd x x ventral neuroblasts  orthodenticle otd x midline  Dichaete D x nervous system  glial cells missing gcm x all glia except midline glia  Netrin-B NetB x midline glia  intermediate neuroblasts defective ind x intermediate neuroblasts  slit sli x called “sli2b” on the tube, midline marker  thisbe ths x x probably intronic  single-minded sim x x mesectoderm, probably intronic  rhomboid rho x x x probably intronic  Neu3 Neu3 x x probably intronic  short gastrulation sog x x x definitely intronic  vein vn x intronic  brinker brk x probably intronic  Dorsal Ectoderm zerknüllt zen x x compare with bee zen!  tailup tup x anterior/posterior axis specification  u-shaped ush x embryonic development via the syncytial blastoderm  Anterior-Posterior patterning Antennapedia AntP X Hox gene, antennal segment  fushi tarazu Ftz X extremely strong, use 0.5 ul  scribbled scrib x polarity of follicular epithelium  Antennapedia noncoding RNA Antp x x x Hox gene, antennal segment  bithoraxoid bxd 3' x x x transcriptional repressor  bithoraxoid bxd 5' x transcriptional repressor  Page 64 of 80 knirps kni x x x trunk segmentation  Kruppel kr x x x gap gene  hunchback hb x x x gap gene  hunchback Hb 5' x x x gap gene  Antp Antennapedia x x x Hox gene, antennal segment  P1 Antp Antennapedia x x x Hox gene, antennal segment  P2 Antp Antennapedia x x x Hox gene, antennal segment  3' Ubx Ultrabithorax x x x Hox gene, segment specification  5' Ubx Ultrabithorax x x x Hox gene, segment specification  3' abdominal A abd-A x sensory organ development  Kruppel Kr 5' x gap gene (probably intronic)  knirps kni X trunk segmentation  wingless wg x x segment polarity  sloppy paired slp1 x head segmental identity  sloppy paired slp2 x head segmental identity  hairy h x x x pair rule  even skipped eve x x x pair rule  tailless tll x x x axis specification  Others lacZ lacZ x x 

Page 65 of 80 AVAILAB LE PROBES FOR TRIBOLIUM I N SIT U HYBRIDISATION

These are shared reagents in labeled boxes at -20ºC Boveri freezer. Ask TAs for help as we have limited quantities. Please make sure you have your entire staining scheme mapped out before you add probes.

PROBE SYMBOL DIG DNP BIO NOTES RATING Mesoderm twist twi x x x mesoderm  short gastrulation sog x mesoderm, vNE  snail sna x x x mesoderm  cactus cac x mesoderm very early  heartless htl x only one FGFR, heartless and breathless  tinman tin x x heart, no early mesoderm expression  Myocyte enhancing factor 2 Mef2 x x late mesoderm  neuralized neur x tom/bearded/neuralized, double with sim  Neuro-Ectoderm rhomboid rho x x highly divergent, mesoderm early  single-minded sim x x ditto, recommend double with twist/snail  slit sli x ventral midline  ventral nervous system defective vnd x x ventral neuroblasts  orthodenticle 1 otd1 x only one of the otds is embryonic. . .  orthodentical 2 otd2 x  Dichaete D x nervous system  Dorsal Ectoderm pannier pan x x amnion, also in the heart  dorso cross doc x  Anterior-Posterior patterning sloppy paired A slpA x x pair rule  sloppy paired B slpB x pair rule  Ultrabithorax Ubx x x Hox gene, segmentation  abdominal A abd-A x x sensory organ development  Abdominal B Abd-B x x sensory organ development  wingless wg x x segment polarity 

Page 66 of 80 AVAILABLE PROBES FOR APIS I N S I T U HYBRIDISATION

These are shared reagents in labeled boxes at -20ºC Boveri freezer. Ask TAs for help as we have limited quantities. Please make sure you have your entire staining scheme mapped out before you add probes.

PROBE SYMBOL DIG DNP BIO NOTES RATING Mesoderm worniu/snail wor/sna x x you find out!  heartless htl x not the best probe, single FGFR  twist twi x early mesoderm marker  tinman tin x heart, no early mesoderm expression  sloppy paired sog x anterior mesoderm, ventral midline  Myocyte enhancing factor 2 Mef2 x late muscle marker  Neuro-Ectoderm simple minded sim x x mesectoderm  Sox Neuro soxN x great probe, try it with fluorescence  slit sli x ventral midline  orthodenticle otd x ventral midline  ventral nervous system defective vnd x ventral neuroblasts  intermediate neuroblasts defective ind x intermediate neuroblasts  glial cells missing gcm x all glia except midline glia  Dorsal Ectoderm zerknüllt zen x extra embryonic, highly divergent from fly 

Page 67 of 80 In situ hybridization for Parhyale Day 1: (3 hours)

1. Rehydrate embryos into PT

1. Fix for 30 minutes in Fix Solution

2. Wash 3 times in PT

3. Wash 3 times in Hyb Buffer at 65 degrees

4. Wash 1 time in Hyb Buffer at 65 degrees for 30 minutes

5. Incubate embryos in Probe Solution at 65 degrees for at least 24 hours (overnight)

Day 2: (6 hours)

6. Remove Probe Solution and save

7. Wash 3 times in Hyb Buffer at 65 degrees

8. Wash 4 times in Hyb Buffer at 65 degrees for 1 hour (4 hours total)

9. Wash 3 times in PT

10. Wash 3 times in PT for 20 minutes (1 hour total)

11. Incubate in Antibody Solution at 4 degrees for 12-18 hours (overnight)

Day 3: (2.5 hours +reaction time)

12. Wash 3 times in PT

13. Wash 3 times in PT for 30 minutes (1.5 hours total)

14. Incubate in 300 ul of BM-Purple for 1 to 24 hours (watch rxn)

15. Wash 3 times in PT

16. Wash 1 time in PT (store in PT until DAPI staining)

17. Wash 1 time in PT with DAPI for 1 hour

18. Wash 1 time in PT Store in 70% Glycerol

Page 68 of 80 FIXED DROSOPHILA EMBRYOS AVAILABLE

These are shared reagents in labeled boxes at -20ºC Boveri freezer. Ask TAs for help as we have limited quantities of some genotypes.

GENOTYPE STAGE NOTES Drosophila - Wild Type wild type (y and w mutations do not affect y w o/n, 2-4, 2-6 development) Drosophila - Mutants and Crosses twist snail 2-4h twist/CyO BSC2381 snail/CyO BSC2311 Tom ~2-4h expanded mesectoderm toll rm9/rm10 ~2-4h also carry dead str.2-Snail TG, don't stain for snail toll rm9/rm10, st2-Nicd 2.5-4.5 also have stripe2 driving activated Notch hsp83-Toll10B-bcd 3' UTR anterior posterior gradient of Toll signaling hsp83-Twist-bcd 3' UTR anterior posterior gradient of Twist tin346/TM3 thank you, Manfred Frasch, no dorsal mesoderm eve[1]/CyO Hypomorph, BSC5344 twi; 24B x DmC15 stain for cardiac Eve, other pericardial markers twi; 24B x TcC15 stain for cardiac Eve, other pericardial markers twi; 24B x TcLB stain for cardiac Eve, other pericardial markers Drosophila - Enhancers (might want to compare to endogenous gene) stripe2-lacZ eve stripe 2 enhancer Tribolium cactus-lacZ large and minimal forms of enhancer in the box compare Dm, Tc, check out AP/DV integration in Drosophila cactus-lacZ Tc Anopheles brk-lacZ this one is in the Atg5 intron! 100 kb away! Dros. sna shadow-lacZ Joung-Woo's paper Dros. sog shadow-lacZ Joung-Woo's paper Other Insects Apis embryos 3 and 4 day collections Tribolium embryos mixed stages, o/n to 5 days

Page 69 of 80 LIVE FLY STOCKS AVAILABLE – MB L 200 9

These are shared stocks on the trolley by the front desk. You can take a tube of the genotype you need back to your bench to work with. If you need help with virgin collection, crossing, embryo collection, dissection etc., as your TAs.

Genotype Abbreviation Notes SPECIES Drosophila melanogaster Wild type Vallecas strain from Madrid UAS LINES UAS-RFP-myr/CyO (1-8) UAS-mRFP see cell outlines in red w; UAS-gma (II) CyO (1-8) UAS-moe-GFP actin-binding protein: see cell outlines UAS-GFP-nls CyO (1-8) UAS-GFP nls see nuclei: first chromosome insert UAS-Ubx IV a 1 (IV a 25) (2, 7) UAS-Ubx-1Va 25 for overexpression of Ultrabithorax protein, homeotic transformations possible UAS-Ubx I a 1 (2, 7) UAS-Ubx-1a 1 for overexpression of Ultrabithorax protein, homeotic transformations possible UAS-Ubx IV a 33/CyO (2, 7) UAS-Ubx-1Va 33 for overexpression of Ultrabithorax protein, homeotic transformations possible UAS-Ubx I a 2 (2, 7) UAS-Ubx-1a 2 for overexpression of Ultrabithorax protein, homeotic transformations possible GAL4 LINES arm-GAL4 CyO (1-8) Arm-GAL4 ubiquitous ectodermal expression en-GAL4 (1, 3) En-GAL4 posterior compartment from embryo to discs nubbin-GAL4 (7) Nubb-GAL4 wing pouch driver A9 GAL4 (wing pouch) (7) Wing-GAL4 A9 wing pouch driver MS1096-GAL4 (7) wing-GAL4 wing driver MS1096 w; NP3537, GAL80(7)1/TM6, Sb Tb (2, 7) Wing- GAL80 on at 18°C -> turns off GAL4 GAL4/GAL80 GAL80 off at 29°C -> GAL4 on MARKER LINES (GFP, YFP, lacZ) w; ubi-DECadherin-GFP CyO (1-8) DE Cad-GFP E-cadherin-GFP driven by an ubiquitious

Page 70 of 80 promoter En-GFP(1, 3, 8) En-GFP posterior compartment & some nervous system His-GFP CyO (1-8) His-GFP histone-GFP under control of an ubiquitous promoter. Moe-GFP CyO (1-8) Moe-GFP ubiquitious squash promoter driving moesin-GFP (actin cytoskeleton) scribbled-GFP CyO (1-8) Scrib-GFP non-muscle myosin GFP w[1118]; P{w[+mC]=GMR-myr-mRFP}7 (6) GMR-RFP RFP enhancer trap w[*]; P{w[+mW.hs]=arm-GFP.P}83 CyO (1-8) Arm-GFP GFP enhancer trap w[*]; P{w[+mC]=His2Av[T:Disc\RFP-mRFP]}III.1 CyO His-RFP 3 RFP enhancer trap; third chromosome (1-8) insert w[*]; P{w[+mC]=His2Av[T:Disc\RFP-mRFP]}II.2 CyO His-RFP 2 RFP enhancer trap; second chromosome (1-8) insert MUTANT LINES bcd[12] p[p]/TM3, Sb[1] (1) bcd loss of function EMS allele cn[1] en[1]/SM5(1, 3) en spontaneous loss of function allele eve[5]/SM6a(1, 3) eve Pair rule y[1] fog[S4]/FM7c(1) fog blocked gastrulation movements ftz[1]/TM3, Sb[1] B54 ts-hypomorph(1) ftz hypomorphic diepoxybutane allele; pair rule phenotype P{ry[+t7.2]=ftz/lacC}4; kni[10] hb[4]/TM3, Sb[1] kni hb gap phenotype; strong amorph EMS allele (1) Kr[2]/CyO, P{ry[+t7.2] = ftz/lacC}1(1) Kr gap phenotype st[1] nos[L7] e[1]/TM3, Sb[1] (1, 8) nos EMS loss of function allele Ubx[bx-3] Ubx[bxd-1] Ubx[pbx-1]/T(2;3)ap[Xa], Ubx triple[1] transformation of haltere to wing ap[Xa] (2, 7) identity Ubx[bx-3] Ubx[Cbx-1] Ubx[1]/T(2;3)ap[Xa], ap[Xa] Ubx triple[2] transformation of haltere to wing (2, 7) identity wg[Sp-1]/SM5(1) wg segment polarity mutant

Page 71 of 80 APPENDIX: EXTRA PROTOCOLS

The following are protocols that you will not be using in this lab, but that you may find useful someday…

Alternative Fluorescent Detection Option: Tyramide Signal Amplification (TSA)

Follow your favorite in situ protocol through the Hyb, hot washes, and PBT rinses. If you are not doing an ISH, but simply an antibody stain, step (1) picks up after the rehydration. Because of the antibodies and TSA kit we have for this course, TSA is possible for antibodies raised in sheep and mouse (in limited numbers) as well as in Rabbit. Be sure to avoid cross reactivity due to biotinylation of probes and TSA reagents.

1 1. Block /2 hour in Roche Western Blocking reagent 1:5 in PBT. 2. Incubate in 1° antibody (e.g. mouse-anti-BIO) diluted at appropriate concentration either O/N at 4°C or 2-3 hrs at room temp. (Pre-absorbed, if necessary.) (if you want to detect signal in other channels by direct labeling, add the appropriate 1° antibodies at this point) 3. Wash 5 times for 10’ in PBT. 4. Re-block 1/2 hour in blocking reagent. 5. Dilute appropriate (i.e. anti-mouse) biotinylated 2° antibody from the Vectastain ABC Elite kit 1:200 into 500 µL of blocking reagent.  Incubate in 2° antibody for 1 hour. (If you want to detect signal in other channels by direct labeling, add the appropriate direct-labeled 2° antibodies at this point). 7. Wash 4-5x in PBT over 1 hr. 8. This is important: 30 minutes prior to use, mix 2.5µL each of solutions A and B from the Vectastain kit in 500µL of PBT. Add 500µL of pre-incubated A+B to the sample for 1 hour. 9. Wash 5-6x in PBT over 45’. 10. Rinse once and rock once for 5’ in TNT buffer. 11. Dilute fluorescent tyramide from NEN/PerkinElmer 1:50 into 300 µL of amp diluent solution (included in NEN kit). 12. Apply fluorophore to the embryos. Remove small aliquot of embryos and let settle into 1mL of PBT after 5’ to check under microscope. (Probably, if it hasn’t stained after 10-15’, or at the outside, 30’, it never will.) 13. IF you are going to do a 2nd color by TSA you have to quench the first HRP reaction before proceeding: inactivate the first peroxidase, by rocking 20’ in PBT + 1% H2O2 - wash 3x PBT, rock 3x PBT 5’ - go back, repeat with a 2nd 1° antibody etc. Note: if doing it a 2nd time, the morphology will suffer… 14. Rinse with 5-6 changes of PBT over 1 hr (more if need be) and mount.

TNT buffer: 0.1M Tris-HCl pH 7.5 0.l5M NaCl 0.05%Tween-20

Page 72 of 80 Heat Fixing Embryos or Imaginal Discs: 1. Make up a stock of salt solution: 4 g NaCl + 300 ml Triton X-100, bring up to 100 ml with water. 2. Dechorinate embryos using bleach in a wire mesh basket. 3. Wash several times with water. 4. Bring small amount of salt solution to boil in a microwave. Use glass scintillation vials for this. 5. Quickly add dechorionated embryos to the solution and immediately add more salt solution and place the vial on ice for at least 5 minutes. 6. Remove all of the salt solution, leaving embryos behind in the vial. 7. Add 5 ml of heptane to the vial and give the embryos a swirl. 8. Add 5 ml of methanol and vigorously shake the embryos for 30 seconds to pop the vitelline membrane off. 9. The popped embryos will sink to the bottom. Collect them and wash them a few times with methanol in an eppendorf tube. 10. Leave the embryos at –20ºC for at least 4 hours. 11. Take embryos out and rehydrate by washing 3x 10 minutes each wash in PT buffer. 12. Now they’re ready for staining!

Page 73 of 80 Clonal Analysis This powerful technique was developed decades ago to study the effects of homozygous mutations on small groups of cells, in genetically mosaic animals. It can be useful for observing the behaviour of wild type and mutant cells during proliferation and morphogenesis, and for analyzing cellular phenotypes of mutations that are embryonic lethal. It was originally done using X-irradiation to induce mitotic recombination at random in somatic cells. When this recombination occurs between sister chromatids of a chromosome carrying a mutant allele in heterozygosis, one of the two daughter cells resulting from the mitotic division will be homozygous for the mutation (instead of both being heterozygous, which would have been the result of a normal mitotic division). Recessive or dominant markers can be recombined onto the chromosomes to allow you to distinguish the progeny of cells in which mitotic recombination has occurred.

Mitotic recombination using X-rays.

Nowadays, mitotic recombination is almost always done using the FRT-FLP system, which has is much more efficient than X rays (20-100% recombinants, compared to 1-10%). 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.

Page 74 of 80

Mitotic recombination using the FRT/FLP system. FLP expression can be driven by a ubiquitous heat shock promoter to obtain clones in all tissues (both soma and germ line), or by tissue specific enhancers to obtain clones in specific tissue types.

Unfortunately in this lab, you will not be able to perform clonal analysis due to time restrictions. But it is a powerful technique that you should be aware of if you are working with flies. Check out the paper on Minute genes and the compartment theory to see how clonal analysis has changed the way we think about, and experiment on, development in Drosophila, and ask Cassandra for more details if you are interested.

L3 wing disc of genotype GFP/+, in which clones of GFP-minus cells have been induced by mitotic recombination. Green: GFP; Red: propidium iodide (nuclei). Clones of cells are identified as GFP-minus cells (arrows).

Page 75 of 80 Xgal staining X-gal staining can be a quick alternative to immunohistochemistry using anti Beta-gal antibodies, or to in situ hybridization using an antisense lacZ probe. Resolution at the cellular level may not be as good with X-gal staining, however, because the blue product produced by the enzymatic reaction can diffuse across cell boundaries if it is present in large amounts. We do not have any flies with X-gal reporters in the lab this year, but below are some examples of X-gal stainings on various Drosophila tissues. When deciding between in situ and X-gal stain, keep in mind that the beta-gal protein is very stable, while the message is very unstable. Thus, for purposes of temporal resolution, in situ is preferred. This protocol assumes you already have fixed, hydrated and washed tissue. Heat fixation will not work for this protocol: use aldehyde fixation only. Tissues should not be treated with Methanol before Xgal staining. Xgal staining works on embryos that have been dechorionated but not devitellinised, because Methanol inactivates the enzymatic activity of the β-Galactosidase enzyme. If you have to devitellinise embryos, you can do it with Ethanol instead of Methanol, but the efficiency will be lower. You could also do it by hand – ask Cassandra for a demo if interested.

Protocol 1. Take 967 ul of Colorisation buffer. 2. Add 33 ul of Xgal solution. 3. Mix and pre-warm at 37ºC for 10 minutes. 4. Incubate tissue in staining buffer at 37°C (water bath or dry incubator are both fine) until you see blue staining. This may take anywhere from 15 minutes to overnight.

X-gal Staining buffer Xgal Solution 62 µl 50 mM potassium ferricyanide 8% Xgal in DMF

62 µl 50mM potassium ferrocyanide 15 µl 20% Triton X-100 30 µl 5M NaCl 1 µl 1M MgCl2 100 µl 10x PBS 697 µl water

Some examples of X-gal stainings:

Cryostat section the adult CNS of a fly, product of the enhancer-trap line C850 crossed to UAS-lacZ, stained with Xgal. The GAL4 line C850 is expressed in the T1 cells of the Early embryo of the genotype optic lobe. ftz:lacZ, stained with Xgal. This strain has a lacZ insert under the control of the ftz promoter, which is expressed in a pair rule pattern. Page 76 of 80 Tribolium castaneum Embryo Fixation (adapted from Dave Kosman) 1. Collect flour, bang out the adults through a coarse sieve. Vortex the remaining flour through a fine sieve (I recommend the Carolina Biological Suppy soil sorting kit, it provides sieves with about the right mesh sizes) to separate eggs from flour. 2. Transfer eggs to a fix basket. Rinse out excess flour and yest 3. Dechorionate the embryos by dunking in 100% bleach for 2-3 minutes (these guys are tough). Periodically squirt some bleach in from the side with a pipet for gentle mixing. 4. Wash the dechorionated embryos thoroughly to remove all traces of bleach. Alternate washes between double distilled (dd)H2O, which causes the embryos to clump together, and the embryo wash buffer, which breaks the clumps. Do a final wash in ddH2O, making the embryos clump, which are then quite easy to be picked up with the wet paint brush. 5. Using the paint brush, transfer the embryos to a scintillation vial containing the following (there are variations on the fix mix, this is what I prefer): • 8 ml 10% ultrapure formaldehyde • 1 ml 10X PBS • 1 ml 0.5M EGTA pH80.0 • 10 ml heptane 6. In general, unlike flies, the more embryos you cram into a fix vial, the better they devitellinize 7. The embryos should now float at the interface between the two phases. Cap the vial and tape it on its side to an orbital platform shaker. Shake it hard for 1 hour (at least) at 220-230 rpm. 8. After shaking, let the bubbles at the interface pop. Completely remove the bottom, aqueous phase with a pipette, avoiding pulling up the embryos. 9. Add about 10-15 ml of cold (-80) methanol, cap the vial and alternately shake and vortex the vial for 5 minutes. The fixed, devitellinized embryos settle to the bottom in methanol. Take an 18 gauge syringe and run the embryos left at the interface through the syringe a couple of times to try to get some more of them to devitellinize. 10. 8.. First remove the heptane, and then the methanol, leaving the embryos at the interface (chances are, many of them are good). Rinse the embryos in fresh methanol until you don’t see any floating crap, just embryos. Gently transfer the embryos to an eppendorf tube using a cut off pipet tip or pipet (you want a large bore, as a small one will pulp them). 11. Embryos can be stored long term in MeOH at –20 C.

1

2 VIII. ANTIBODY LIST FOR 2009 (+OTHER STAINS) Where is it? Rating

Concentration in 300 µl PT+N Primary Antibodies Staining Pattern Species Species Specificity / Cross-reactivity / Comments (assumes overnight incubation)

Anti-Pax3/7 (pair-rule, segment polarity, neural gene family) In Drosophila, pair- 4°C Boveri DP312 rule pattern early (prd), then segmental pattern (gsb), then CNS pattern (gsb- mouse mAb 1 : 30 All animals tested including arthropods, annelids,and vertebrates  Fridge neuro)

Anti-even-skipped (pair-rule gene). In Drosophila, pair-rule pattern early, subset Drosophila, Tribolium, most insects (3B9 better for grasshoppers), some crustaceans 4°C Boveri 2B8 mouse mAb 1 : 40  of CNS neurons and subset of dorsal muscle and heart later (mysids) Fridge

Almost all insects (not lepidoptera), most crustaceans (including mysids, but not Anti-engrailed/invected (segment polarity gene). In Drosophila, segmental pattern 4°C Boveri 4D9 mouse mAb 1 : 40 Artemia and Triops), many vertebrates including zebrafish and chickens (not mice,  early, CNS pattern later Fridge weak in Xenopus)

Almost all insects (including lepidoptera), some crustaceans (including Artemia and Anti-engrailed/invected (segment polarity gene). In Drosophila, segmental pattern 4°C Boveri 4F11 mouse mAb 1 : 30 Triops), no vertebrates. Superior to 4D9 for Drosophila imaginal disc staining, but  early, CNS pattern later Fridge not as good for embryos

4°C Boveri FP3.3 Anti-Ubx (homeotic gene). In Drosophila, regional expression pattern mouse mAb 1 : 20 Drosophila specific  Fridge

Stains nuclei of all neurons (elav gene product, encodes neural-specific splicing 4°C Boveri 9F8 mouse mAb 1 : 10 Drosophila specific  factor) Fridge

4°C Boveri BP102 Stains all CNS axons (unknown antigen) mouse mAb 1 : 200 Drosophila specific  Fridge

Stains cytoplasm and cell membranes and axon of all PNS neurons plus a subset 4°C Boveri 22C10 mouse mAb 1 : 25 Drosophila specific  of CNS neuron Fridge

Anti-hunchback (gap gene). In Drosophila, gap pattern early, CNS and other 4°C Boveri 1G10 mouse mAb 1 : 10 Drosophila specific  tissues later. Fridge

Anti-hunchback (gap gene). In grasshoppers, gap pattern early, CNS and other 4°C Boveri 7C11 mouse mAb 1 : 10 Grasshopper specific  tissues later. Fridge

Anti-even-skipped ("pair-rule gene) In grasshoppers, posterior domain early, CNS 4°C Boveri 3B9 mouse mAb 1 : 20 Grasshopper specific (better than 2B8 for grasshoppers)  + heart later Fridge

Grasshopper (but 3B9 better for grasshopper), some additional insects (but not 4°C Boveri 7H5 Anti-even-skipped ("pair-rule" orthologs) mouse mAb 1 : 10  Drosophila), some crustaceans (works fine in Artemia, inconsistently in Triops) Fridge

4°C Boveri FP6.87 Anti-Ubx/abd-A (homeotic genes). In Drosophila, regional pattern mouse mAb 1 : 20 All arthropods VERY LIMITED QUANTITY AVAILABLE - SAVE AND REUSE  Fridge

4°C Boveri 8C11 Anti-Antp (homeotic gene) , regional expression mouse mAb 1 : 20 Many insect species including Drosophila, Tribolium, and grasshoppers  Fridge

4°C Boveri 1D11 (ask Nipam) Anti-Scr (homeotic gene), regional pattern mouse mAb 1 : 300 Drosophila specific  Fridge

4°C Boveri 1D4 Anti-Drosophila fasII (subset of CNS axons) mouse mAb 1:30 Drosophila specific  Fridge

4°C Boveri 7G10 Anti-Drosophila fasIII (subset of CNS axons) mouse mAb 1:20 early, 1:30 late Drosophila specific  Fridge

Page 1 4°C Boveri 3B11 Anti-Grasshopper fasI (subset of CNS axons) mouse mAb 1:5 Grasshopper specific  Fridge

4°C Boveri 8C6 Anti-Grasshopper fasII (subset of CNS axons) mouse mAb 1:5 Grasshopper specific  Fridge

4°C Boveri 6F8 Anti-Grasshopper fasIV (subset of CNS axons) mouse mAb 1:5 Grasshopper specific  Fridge

4°C Boveri ADL84.12 Anti-lamin DmO Mouse IgG1 1:10 Drosophila  Fridge

4°C Boveri Anti-vasa Anti-vasa Rat IgM 1:20  Fridge

4°C Boveri 13C9 Anti-Robo Mouse IgG1 1:20 Drosophila  Fridge

4°C Boveri 15H2 Anti-Robo3 cytoplasmic Mouse IgG1 1:20 Drosophila  Fridge

4°C Boveri Anti-Antp 8C11 Anti-Antennapedia Mouse IgG 1:20 Drosophila  Fridge

4°C Boveri Anti-AbdB (1A2E9) Anti-Abdominal-B Mouse IgG1 1:50 Drosophila  Fridge

4°C Boveri 8D12 Anti-Repo Mouse IgG 1:20 Drosophila  Fridge

4°C Boveri Single minded Anti-single minded Mouse IgG1 1:10 Drosophila  Fridge

4°C Boveri ab50567 Rat monoclonal (MAC141) to Tropomyosin Rat mAb 1:100 Crossreactive  Fridge

4°C Boveri T6793-.2ML Anti-acetylated tubulin (clone 6-11B-1, ascites fluid) mouse mAb see Nipam Crossreactive  Fridge

4°C Boveri C2206-.2ML Anti-beta-catenin Rabbit see Nipam Crossreactive  Fridge

4°C Boveri 51.4H9 Anti-Islet-2 (mouse IgG1; broadly crossreactive) see Nipam Crossreactive  Fridge

4°C Boveri E7 Anti-tubulin (beta-) mouse IgG1 see Nipam Crossreactive  Fridge

4°C Boveri 06-570 Anti-phospho-Histone H3 (Ser10) Mitosis Marker; 1 mg/ml (polyclonal) Rabbit 1:100 Crossreactive  Fridge

Anti-Pax3/7 (pair-rule, segment polarity, neural gene family + repo) In 4°C Boveri DP311 mouse mAb 1:30 All animals tested including arthropods, annelids,and vertebrates  Drosophila, pair-rule pattern early, then segmental pattern, then CNS pattern Fridge

4°C Boveri Anti-PhEve Anti-even-skipped in Parhyale ("pair-rule" ortholog) RAT 1:500 Parhyale-specific  Fridge

Page 2 Stains nuclei of all neurons (elav gene product, encodes neural-specific splicing 4°C Boveri 7E8 RAT mAb 1:10 Drosophila specific  factor) Fridge

Anti-even-skipped (pair-rule gene) In Drosophila, pair-rule stripes early, CNS + 4°C Boveri 10900 (M. Biggin) RABBIT 1:100 Drosophila, other species not tested  heart later Fridge

4°C Boveri 11175 (M. Biggin) Anti-fushi tarazu (pair-rule gene) In Drosophila, pair-rule stripes early, CNS later RABBIT 1:125 Drosophila, other species not tested  Fridge

4°C Boveri 11279 (M. Biggin) Anti-Ubx (homeotic gene) , regional expression RABBIT 1:50 Drosophila, other species not tested  Fridge

4°C Boveri 20254 (M. Biggin) Anti-snail (mesoderm) RABBIT 1:50 Drosophila, other species not tested  Fridge

4°C Boveri 20265 (M. Biggin) Anti-twist (mesoderm and muscle) RABBIT 1:100 Drosophila, other species not tested  Fridge

4°C Boveri 20405 (M. Biggin) Anti-teashirt (modifier of Hox gene function). Expressed in trunk region. RABBIT 1:125 Drosophila, other species not tested  Fridge

4°C Boveri Anti-pHistone Anti-phospho histone (present on chromatin during mitosis) RABBIT 1:200 All animals tested  Fridge

4°C Boveri Anti-Spalt Readout of dpp signaling in imaginal discs. In Butterfly discs, marks eyespot RABBIT see Nipam Drosophila embryos and discs, Butterfly discs  Fridge

4°C Boveri Anti-DsRed Anti-DsRed (red fluorescent protein) RABBIT 1:3000 n/a  Fridge

goat 4°C Boveri Anti-HRP Alk Phos conjugated Stains all neurons (binds carbohydrate epitope present on many neural proteins) see Nipam All arthropods tested, most all other Ecdysozoa as well  No 2°Ab needed Fridge

goat 4°C Boveri Anti-HRP FITC conjugated Stains all neurons (binds carbohydrate epitope present on many neural proteins) 1:200 All arthropods tested, most all other Ecdysozoa as well  No 2°Ab needed Fridge

goat 4°C Boveri Anti-HRP RITC conjugated Stains all neurons (binds carbohydrate epitope present on many neural proteins) 1:300 All arthropods tested, most all other Ecdysozoa as well  No 2°Ab needed Fridge

4°C Boveri 1B1 Adducin Mouse 1:100 Fusome component  Fridge

4°C Boveri 1B11 Bicaudal D Mouse 1:20 In embryos, pole cells up to S9-10; germaria and oocytes  Fridge

4°C Boveri 1D12 Gurken extracellular domain Mouse 1:30 Anterodorsal position in oocyte  Fridge

4°C Boveri 3A9 Alpha spectrin Mouse 1:10 Fusome component  Fridge

4°C Boveri 4C2 Bicaudal D Mouse 1:30 In embryos, pole cells up to S9-10; germaria and oocytes  Fridge

4°C Boveri Anti-Gro Groucho Mouse 1:5 Translational repressor  Fridge

Page 3 -20ºC Boveri Beta-gal beta-galactosidase Rabbit 1:5000 Can be used with transgenic lacZ lines  freezer

4°C Boveri C17.9C6 Notch intracellular domain (NICD) Mouse 1:40 Transmemebrane receptor - intracellular domain only  Fridge

4°C Boveri C458.2H Notch extracellular domain (NECD) Mouse 1:40 Transmembrane repector – extracellular domain only  Fridge

4°C Boveri C594.9B Delta extracellular domain Mouse 1:4000 Various aspects of nervous system throughout development  Fridge

4°C Boveri Cmnb-1 Crumbs Mouse 1:10 approx. Apical membranes of epithelial cells  Fridge

4°C Boveri Cq4 Crumbs (aa 737-1703) Mouse 1:5 Apical membranes of epithelial cells  Fridge

4°C Boveri DCAD-2[1] Anti E-cadherin Rat 1:15 Adherens junctions  Fridge

4°C Boveri DN-Ex#8[2] Anti N-cadherin Rat 1:20 Adherens junctions  Fridge

Anterior localization in oocytes S8-10; ubiquitous in oocytes before and after that 4°C Boveri Orb 4H8 Orb Mouse 1:30  stage Fridge

Other Dyes Description (fluorescent color) Species vol. in µl/300 µl PT+N Species Specificity / Cross-reactivity / Comments

DNA Dyes

4°C Boveri DAPI (Blue) n/a 1 µg/ml labels DNA (nuclei), detect as blue flourescence  Fridge

4°C Boveri Propidium iodide (Red) n/a 1:500 labels DNA (nuclei), detect as red flourescence  Fridge

Polymerized Actin Dyes

4°C Boveri Phalloidin 488 (Green) n/a 1:500 labels muscles (muscles are made of actin)  Fridge

4°C Boveri Phalloidin 555 (Red) n/a 1:500 labels muscles (muscles are made of actin)  Fridge

labels muscles (muscles are made of actin) 4°C Boveri Phalloidin 647 (Far Red) n/a 1:500  CONFOCAL ONLY!!! Fridge

Page 4 4°C Boveri DIG Digoxygenin Sheep for in situs: detect probes 1:400  Fridge

4°C Boveri DIG Digoxygenin Mouse for in situs: detect probes 1:400  Fridge

for in situs: detect probes – 4°C Boveri DIG-AP Digoxygenin Mouse 1:400  Alkaline Phosphate Conjugated Fridge

4°C Boveri DNP Dinitrophenyl-KLH Rabbit 1:400 For in situs – detect probes  Fridge

sheep Anti-Fluorescein Anti-Flourescene (green fluorescent dye) 1:4,000 React with BCIP/NBT No 2°Ab needed

4°C Boveri BIO Biotin Mouse 1:400 For in situs – detect probes  Fridge

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