Exercise 6 RNAi screening in Drosophila: Dendrite morphology Learning Objectives: skills acquired: You will learn how Drosophila is used for RNAi knockdown of candidate genes, how cell type-specific expression of RNAi hairpins and fluorescent proteins is performed, very basic Drosophila genetics (setting up a cross and collecting progeny), and imaging of neurons in whole, living animals. concepts put into practice: You will learn how to interpret data based on comparison with a control. You will learn to generate a hypothesis based on observation and develop an analysis method to test your hypothesis. In other words, you will be doing a real experiment that no one else has ever done before! Overview of the Experiment (see also the background section at the end): KNOW FOR QUIZ For this experiment to be successful you need to understand what you are doing and why you are doing it. If something is not making sense, please ask as we go along. Overall goal: In this experiment, you will be identifying genes required to control two aspects of dendrite organization: 1. dendrite shape 2. localization of proteins to specific regions within dendrites Why is this important: Dendrites are the receiving end of neurons- they receive signals either from the outside world (as in the sensory neurons you will be looking at) or from other neurons. These signals are then integrated and, based on the structure of dendrites and amount and frequency of signal, an action potential may be sent or not sent. The action potential propagates long distances down the axon and then causes release of synaptic vesicles. Neurotransmitters released by the synaptic vesicles are sensed by surrounding cells and the signal has moved to its next processing destination. Dendrite shape is a key part of how signals are received and processed by neurons. First of all, the long branched shape of dendrites allows them to reach out to the cells or other sources of the signals they receive, and second the specific shape determines how electrical signals move through the dendrites and how they are integrated over space and time. Regional specialization of dendrites (including localization of proteins and organelles to different spots) most likely allows dendrites to take on their specific shapes, and for different regions of the dendrite arbor to receive and process signals differently (note that the significance of localization of organelles to branch points is a hypothesis- to test this hypothesis we would need to mess up the organelle localization and then see what went wrong). Approach you will take: You will be examining two different fluorescent markers in a sensory neuron in Drosophila larvae. One of these markers (Ank2-GFP) is part of the Ankyrin-2 scaffolding protein. This protein often functions in axons to target channels to the place where the action potential initiates. As you will see, this GFP-tagged protein is highly concentrated at dendrite branch points and functions to localize other proteins there. The other marker (mCD8-RFP) outlines the whole cell. You will express a hairpin RNA to knock down a specific gene in this cell and then determine whether the shape of the cell or localization pattern of Ank2-GFP is different from control cells. Each person in the class will be targeting a different gene and so as a group you will test about 20 candidates and be able to determine which ones play a role in controlling development of dendrite shape and protein localization. Once we know which genes and proteins are involved with determining dendrite shape and Ank2 localization we can start to come up with hypotheses about how they might work to control dendrite development. ** if you are unfamiliar with any of the terms being used, like GFP or RFP, please look them up! There is a pretty fun entry on GFP (green fluorescent protein) on Wikipedia. RFP stands for red fluorescent protein. Can you figure out roughly what wavelengths you will use to excite GFP and RFP and what wavelengths they will emit?

Important note: this is a little different from previous experiments in that you will be responsible for all the steps of the experiment once it begins. Also, each day here is a consecutive day, not a lab period, so you will need to come in and do something pretty much every day of the week. If you are not taking care of your flies EVERY DAY (except the weekend), then you will not be able to do the experiment. Please be responsible and email me (mur22@psu.edu) if you are not sure you know what you are supposed to be doing on a particular day. I am happy to help- either by email or by popping over to the lab room to meet you. Procedure: Day one (Friday, week 1=Oct 3): starting flies on a virgining cycle. Introduction to Drosophila genetics and RNAi. Materials Drosophila test stock: UAS-Dicer2, UAS-mCD8-RFP/CyO; 221-Gal4, UAS-Ank2-GFP/TM6 Bottles and vials of Drosophila media practice flies- for identification of males and females Procedure Part 1: getting your flies ready to virgin 1. Label fresh bottle of media with full genotype of test stock and your name 2. Remove adult flies from your test stock to a fresh bottle of media- this will be your stock of flies; when new adults start to emerge in 10-14 days, you will switch to this for your source of virgin females 3. The bottle with pupal cases that you just removed adults from will be the source of your virgin flies for the first part of the experiment 4. Place the bottle of flies that now has no adults at 18C or 25C over the weekend- if the flies are not quite ready to virgin you will choose 25C to get them growing faster so they will produce adults Monday, if there are already dark pupal cases that look like they have mini adult flies in them, put your bottle at 18C **NOTE** we will start on a Friday, so the flies will be sitting at 18C or 25C all weekend. This means that the newly emerged flies will NOT be virgins on Monday- you will just need to dump adults and collect females less than 8h later. Part 2: practice identifying and collecting females 1. Take a bottle of practice flies to one of the four fly genetics stations 2. Turn on CO2 by turning handle directly on the top of the tank- DO NOT adjust any of the dials on the regulator; it should already be set up right. The CO2 gas will bubble through a flask of water- this reduces static, and also reminds you that the gas is on. It will then reach a porous pad and form a pillow of CO2 near the top of the pad. This stops the flies moving when you place them on the pad. 3. Turn on light source for dissecting scope. 4. Invert bottle, flies will move away from the stopper. 5. When few flies are near the stopper, open it while keeping the bottle inverted. 6. Tap neck of bottle on fly pad, and hold bottle there for a few seconds to stop tons of flies getting away. 7. After you have some flies on the pad to look through, keep bottle inverted while replacing stopper. 8. Separate flies into piles of males and females by gently moving them on the pad with a paintbrush. 9. Ask instructor to check you can reliably separate males and females: you need to be able to do this on your own after this session! **safety note- please turn on CO2 slowly. This is a pressurized gas and can cause the stopper to pop out of the flask. Also be familiar with regulator parts- look at the picture on the tank.

Overview of virgining flies KNOW FOR QUIZ

There are several different methods to collect unmated or virgin females. You need unmated females because once they have mated they can store the sperm for a long time, and you want to set up a cross with a male that has a specific genotype. We will use a time-based method to collect virgins. After adults emerge from the pupal case, it takes them about 8h at room temperature, or 18h at 18C (speed of development depends on temperature), to become sexually mature. This means that females younger than 8h at room temp (or 18h at 18C) are virgins. So if you clear a bottle of adults, leave it at room temp and come back within 8 hours all the new females you find in the bottle when you return will be virgins. If you always come to the lab within 8h when you have your flies at room temp, or within 18h when they are at 18C, all the females will be virgins, and you will quickly have enough to set up a cross. If you miss a time point, you will not know which of the flies in your bottle are virgins and so you should throw them all out and start on your virgining cycle again. Day two (Monday, week 2=Oct 6): Making sure everyone is good with fly genetics and experiment overview, and virgining your test stock Materials Vials of Drosophila media Procedure Part 1: collecting females 1. Return to the lab Monday MORNING 2. Remove all adults from your bottle that was at 18C or 25C over the weekend. (Discard the adults in fly morgue) 3. Come to class, we will have an overview lecture, then virgin your flies: 4. Take your bottle to one of the fly genetics stations, turn on light and CO2. 5. Invert bottle, remove stopper and bang out ALL flies on the CO2 pad. 6. Visually inspect bottle to ensure all adults are out. 7. Collect virgin females, and put in labeled food vial. 8. Put flies in 18C incubator overnight. Part 2: setting up a cross (do in class time) 1. (during lab period). You may now have enough female flies to set up the first cross (if you dont today, you will by tomorrow afternoon)!! 2. Place 10-30 female flies in a single vial (if you only have a few flies, work with these then add more as you collect them over the next couple of days). 3. Collect about 10 male flies from our control RNAi stock. Notes about the control: everyone in the lab will use the same control, BL25271, which targets gamma-tubulin 37C. This gene is expressed only in the early embryo and not somatic cells including neurons. Also, we have never observed any phenotypes from expressing this RNAi. This is one possible control. I am not sure it is the absolutely ideal one, but this is something that I would like you to think about (see also Background, RNAi in Drosophila section). 4. Add female tester line flies and male RNAi flies to bottle with air holes, bang flies down in bottle to stop them escaping, cover with food cap, and secure food cap with tape. 5. Label your collection bottle!! 6. Leave at room temperature overnight.

Day three (Tuesday, week 2= Oct 7): continued virgining and collecting embryos Materials Vials of Drosophila media, food caps Procedure 1. Come in twice to collect virgin flies. If you did not have that many female flies (less than 20) in the cross you set up yesterday, add your new females into the collection bottle. 2. Change food cap one of the times you come in to virgin. Discard (scrape out) food in cap, put empty cap in cleaning tub. The flies usually do not lay many embryos their first day of collection. Overview of collecting embryos for the RNAi experiment KNOW FOR QUIZ Females of the tester line will mate with males that contain an RNAi transgene. Embryos will contain all the transgenes from the mother and the transgene that encodes a double-stranded RNA from the father (how many copies of each will the progeny have? will all progeny have the same genotype?). We will therefore be able to analyze dendrite shape in neurons in which a particular RNA has been targeted for destruction (do we know all the RNA is gone? Do we know all the protein is gone?). We will let the females lay eggs (embryos) on the food cap for roughly 24h. Then we will take that food cap and age it for 2 days at 25C, so we get larvae in which we can analyze dendrite shape. When you take off a food cap you will replace it with a fresh one. The idea is to change the cap every day and start aging the animals so that you have larvae to image each day down the road. You may not be able to come in and take pictures each day, but whenever you can, you will have animals ready. We typically set up experiments this way because it is quick to collect embryos, and not so quick to image, so you want to be ready to image whenever possible and not be limited by lack of animals. This will be the philosophy for the rest of the experiment. Also, you want all the animals to be about the same age, so you can only image ones that have had a food cap on for one day (not two days- if you forget to change the cap, throw it out and start collecting again). Days 4, 5 and 6 (Wednesday-Friday, week 2=Oct 8 - 10). Virgining flies and collecting embryos. Materials Vials of Drosophila media, food caps, 10 cm petri dishes (empty) Procedure for each day 1. Come in twice to collect virgin flies. The second time, leave any spare virgins and your bottle in the 18C incubator. 2. Change food cap. Clean up old food caps! The first embryos ones we will keep to image are the ones I will collect Saturday. I will put them into a 10 cm petri dish. And will label with genotype, your name, and when it was collected. I will then put them in the 25C incubator. We will age collected caps for 2 days before imaging- so the ones I collect on Saturday will be ready to look at Monday, the ones I collect Sunday will be ready for Tuesday, and the ones you collect Monday will be ready for Wednesday Day 6 (Friday, Week 2=Oct 10) QUIZ Virgining flies, collecting embryos and learning to mount and image. In class we will talk about good science.

We will also begin to learn how to collect images of larvae (see instructions below for Day 9). It will be a zoo if everyone tries to learn on the same day, so we will need a plan! Day 7 and 8 (Sat and Sun, week 1=Oct 11 and 12). Collecting embryos. Materials Food caps, 10 cm petri dishes (empty) Procedure 1. Instructor will change all the caps from the class each day and keep them so you can image next week! Day 9 (Monday, Week 3=Oct 13) Virgining flies, collecting embryos and learning to mount and image. Materials Vials of Drosophila media, food caps, 10 cm petri dishes (empty), microscope slides with agarose, 22X40mm coverslips, tape, fine forceps, 35 mm dish, pipettor Procedure 1 (outside lab period). Come in twice to collect virgin flies. 2. Change food cap (dont forget to keep cap with embryos on it! Put in clearly labeled 10 cm dish- name, genotype, date, and store in the 25C incubator) 1 (during lab period). Find your cap with larvae (from Saturday). 2. Mount a single larva on agarose slide using live imaging protocol at the end of this document. It may take some practice to be able to mount a larva such that it holds still, but is still alive. 3. On the fluorescence microscope, find the ddaE cell under the polychroic filter. Notes on filter set: this polychroic filter allows both blue and green light to hit your sample. That means that both the GFP and RFP will be excited. Both emission wavelengths (green and red) can then pass up through the filter set to your eyes/camera. 4. Take pictures that cover the entire comb dendrite- we will need to reconstruct these into a complete image of this dendrite afterwards. You will need to be efficient at imaging to avoid bleaching the cell before you get a complete picture. Some imaging tips: use manual exposure between 100 and 500ms, use the neutral density filter (slider in front of shutter) to reduce light on sample, and thus also reduce bleaching, close shutter whenever you can, you can use the freeze function instead of taking a high res picture this will also make for files that are easier to handle later, about 1000 x 1000 pixels is plenty of resolution. Remember to save your images (tif format preferred). To ensure that we can compare images from everyone we may want to come up with additional guidelines for imaging. 5. Take images of one ddaE comb dendrite per animal, then go get a new larva. We want to do this in case RNAi is variable- we would like to sample from several animals, instead of getting all our data from only a few. 6. Please reuse your agarose slide (but not the coverslip). Just lift off coverslip (remember to put in glass waste) and pick off larva with forceps. Then the slide is ready to go again with the next larva. Important note: everyone in the class needs to be imaging ddaE cells in the same segment so that we can compare everyones data. To identify the right cell: using the 10X objective find the larvas nose. Work your way back from there. We will use the second set of neurons behind the nose- they will be on either side of the back end of fluorescence from the ventral ganglion.

Tuesday-Thurs, week 3=10/14-10/16- keep virgining and changing caps so you make sure to have animals to image This week you will be collecting images for your control cross. Take 15 good images of 2-day larvae from the control cross!! You need to get the hang of imaging. If you try it on your own and have questions or need help then get in touch with me (mur22@psu.edu). I can meet you and help you. It is critical that everyone take good images! At some point during the week you will want to update your flies: 1. Discard first cross (its getting kind of old): freeze flies, wash bottle. 2. Set up new control cross with tester virgins you have been collecting, and VDRC# 33320, Rtnl2 males. Use this cross to collect embryos for the next week. 3. Also, if the bottle you are virgining from is getting old, switch to a new bottle (transfer adults into a new bottle so you have a backup stock). Friday, week 3=10/17. Virgining flies, collecting embryos, setting up experimental cross and assembling images. Materials Vials of Drosophila media, food caps, 10 cm petri dishes (empty Procedure 1 (outside lab period). Come in twice to collect virgin flies. 2. Change food cap. 1 (during lab period). Set up new cross with experimental RNAi line. Each group will get a different RNAi line from VDRC. 2. I encourage you to keep your control cross going over the weekend so that you can collect more control data after we go through it all on Monday- some of your images may not be quite what the class is looking for. If you need to take better control images, you can do this next week. 3. I will show you how to find out what gene you are targeting (VDRC web site) and how to find out what is already known about this gene (flybase and pubmed). 4. We will go through assembling the data in class, and I will show you examples. Your control data set needs to be posted on our class Angel site BY SUNDAY NOON (10/19) This should be a power point file with 15 GOOD images of the ddaE cell in the second hemisegment back. You will be graded based on handing this in on time, and also quality of images- including whether they are of the correct cell. However, I will modify your grade if you improve your data set after we go through it together in class Monday. Really the goal is to make sure everyone has usable control and experimental data sets. Sat-Sun = 10/18 and 10/19 Instructor will collect embryos. Mon-Fri week 4= 10/20-10/24. Collecting embryos, finishing control data and starting to collect experimental data.

Materials Food caps, 10 cm petri dishes (empty), microscope slides with agarose, 22X40mm coverslips, tape, fine forceps, Schneiders media, 35 mm dish, pipettor Procedure 1. change cap each day on your experimental cross 2. finish collecting images for your controls 3. by the end of the week you should start to have larvae to image from your experimental cross. In class Monday: we will evaluate control data from the class to make sure everyone is on track. Sat and Sun, week 4=10/25-10/26. Collecting embryos. Materials Food caps, 10 cm petri dishes (empty) Procedure 1. Instructor will change all the caps from the class each day. Mon-Thurs week 5=10/27-10/31. Collecting embryos, finishing experimental data. Materials Food caps, 10 cm petri dishes (empty), microscope slides with agarose, 22X40mm coverslips, tape, fine forceps, Schneiders media, 35 mm dish, pipettor Procedure 1. change cap on Monday- Wednesday on your experimental cross- this means last day for imaging will be Friday!! 2. collect images for your experimental neurons 3. FINISH collecting data by Friday 4. look at images as you go to make sure you can see the whole comb dendrite to make sure you are on track to post your data by Friday evening. In class Monday Before class read dendrite screen paper. We will discuss the paper in class- this is to help prepare you for your data analysis next week. Friday, week 5=10/31. Post data by 9 pm. 10 of the 30 points of your grade in the final lab report will be based on the presence of complete data by 9 pm!!! If it is not there at this time, you will be assigned a 0 for this part. Procedure 1. post your control and experimental images in the dropbox on Angel. 2. Each image of a comb dendrite should be a single page in a powerpoint document. Make one document for your 15 control images and one document for your 15 experimental images. Label each with your last names. Put the VDRC number on the first page. Days 27-28 (Sat and Sun, week 4= 11/01 and 11/02). Look through all data and generate one or more hypotheses.

Materials Your brain + computer + images Procedure 1. look through your control images and sets of controls from several other groups to get a sense of what normal for this dendrite is: keep and eye on both shape and mito-GFP position 2. look through each of the experimental data sets and pick out ones you think might have a phenotype 3. for each data set you think has a phenotype generate a simple hypothesis about what is different than the control. Some examples of hypotheses are: -when we reduce levels of protein Y, fewer dendrite branches from the main comb trunk are present. -when we reduce levels of protein X, dendrites are shorter. -when we reduce levels of protein Z, mito-GFP is no longer targeted specifically to branch points. 4. come up with an idea of how to test your hypothesis based on the data. For the examples above: -count number of branch points off main trunk in control and Y experimental neurons to determine if there really are fewer when levels of Y are reduced. -measure overall dendrite length in control and X neurons to determine whether they are shorter in X. -count number of branch points with clear mito-GFP puncta and total number of branch points, compare ratio of occupied/total branch points in Z RNAi and control cells Note: for some of the quantitation to work, we all need to be assembling our images at the same magnification and putting them into powerpoint at the same size. The magnification is not a problem- we are all using the same scopes and same objective. Please put the images into the powerpoint file such that the longest dimension is 8 inches. Monday, week 6=11/03. Discuss hypotheses in class. Procedure 1. come to class and go through your hypothesis with me to make sure you are on track- this session is to help you with the report. The better prepared you are, the more helpful this session will be. I will be there for you to bounce ideas off of and talk through the data. But I will be relying on you to come up with the agenda. It is in your best interest to make use of me in this class period! Rest of week 6. Perform data analysis, write report and prepare to present your hypothesis and data analysis to the class on Friday! Report is due 11/7 before class. In class on 11/7 each student will present their hypothesis and analysis to the class in a 5-10 min powerpoint presentation. Monday, Week 7= 11/10 We will finish our discussion of the class results, and also develop ways to build upon this data in future studies. This session will also serve as review for the exam. Friday, Week 7= 11/14 Exam on Exercise 6- RNAi in Drosophila Background- KNOW FOR QUIZ Drosophila life cycle Embryos are deposited on the food surface by the mother. The embryo starts off with most cells

roughly equivalent and within 24 hours the body plan is laid out and the embryo is ready to hatch into a larva. The job of the larva is to eat and grow. As they do this they go through several molts. Larval development takes 5-6 days. At that point the animal is large enough, and crawls away from its food, secretes a hard shell around itself (pupal case) and starts to digest away its larval structures- muscles etc. The adult structures are then built from pockets of tissue called imaginal disks that were set aside during embryogenesis and grew in size during larval life. After 5-6 days a complete fly is present inside the case and it breaks out and expands its wings. After about 8 hours it is ready to mate and start the whole process over again. Basic Drosophila genetics Drosophila genetics is incredibly well-developed and there are phenomenal tools available. For your experiments you will not need most of them, but a quick intro will probably be helpful. There are 4 chromosomes in Drosophila: X/Y; 2; 3; 4 (in genetic notation chromosomes are separated by semicolons). Females have 2 X chromosomes, males have an X and Y; Y is small, but has a few key male-related genes. The fourth chromosome is very small and so doesnt show up in genetics much. In the crosses you will be doing all the transgenes are on the second or third chromosome, which means it doesnt matter which comes from the mother or father. Of course Drosophila is a diploid and so has two copies of each chromosome, unless it is a male which has one X and one Y. If a mutation or transgene is homozygous it is present on both copies. If it is heterozygous it is only present on one copy.

Four chromosomes of Drosophila, indicating relative lengths. You can see that 1/5 of the genetic material is on X and each of the two arms of 2 and 3, while 4 is very short. Balancer chromosomes are one of the fabulous tools available for Drosophila genetics. They allow heterozygous mutations/ transgenes to be maintained stably through many generations. Please see the intro to balancer chromosomes in wikipedia: http://en.wikipedia.org/wiki/Balancer_chromosome We will be using one balancer chromosome in this lab. It is a third chromosome balancer and will show up in your tester stock to varying extents. Remember the tester stock genotype is: UAS-

Drosophila life cycle.

Dicer2, UAS-mCD8-RFP/CyO; UAS-mito-GFP, 221-Gal4/ TM6. TM6 is the balancer chromosome. Because it is a balancer, the 221-Gal4 and UAS-Apc2-GFP will never be able to recombine away from one another, and TM6 also has useful markers so we can tell which animals the are produced when we cross this line to another one (for example an RNAi) have the TM6 chromosome and which have the UAS-mCD8-GFP, 221-Gal4 chromosome. TM6 has a larval marker, Tb (tubby), which makes the larvae short and fat. So When you cross your tester strain to an RNAi strain you want to select against Tubby larvae so that you have the GFP marker to look at. Bottom line: select against short fat larvae when you choose which ones to mount and image! CyO is also a balancer chromosome, and, like TM6, suppresses recombination. Its visible marker is Cy, or curly, which means that flies with this chromosome have curly wings. Some of the flies in your bottle will have the CyO chromosome, and others will be UAS-Dicer2, UAS-mCD8-RFP homozygotes. If you have lots of virgins to choose from, will you get higher numbers of useful larvae if you use the curly winged females or straight winged females? Unlike TM6, CyO does not have a marker that is visible in larvae, so you will not be able to tell which animals have this chromosome before you mount them on a slide for imaging. If the animals do have this chromosome, what will you see when you get to the fluorescence microscope? Binary expression system for Drosophila We will be expressing large hairpin RNAs, dicer2, and GFP- and RFP- tagged proteins using the Gal4-UAS binary expression system. This system was developed for Drosophila by Andrea Brand in Norbert Perrimons lab: http://www.ncbi.nlm.nih.gov/pubmed/8223268 The coding sequence of the yeast transcription factor, GAL4, is inserted downstream of an enhancer that turns on in a particular set of Drosophila cells, in our case in a couple of sensory neurons. The GAL4 transcription factor is then made in these cells. It does not bind to any native Drosophila promoters and so just sits in the cell not doing much unless another transgene is present which contains the binding site for GAL4 (the binding site is named UAS for upstream activating sequence. If you put a gene of interest next to the UAS sequence, then it will be controlled by the GAL4 transcription factor, so it will be turned on in whichever cells the GAL4 comes on. The advantage of this binary system is that you can mix and match GAL4 drivers with UAS constructs. So, for example, in our experiment we can express a membrane marker in a few peripheral neurons with the 221 GAL4 driver. We will also express mito-GFP to track its localization,

From Dow, 2007 JEB. Diagram of the Gal4-UAS expression system in Drosophila. In this case flies containing a Gal4 driver are crossed with flies containing a UAS-driven reporter. In our case we have several more transgenes present in one parent, but the overall idea is the same.

and in similar experiments we could express markers that would allow us to track position of mitochondria, microtubule behavior, membrane trafficking etc. In order for this system to work, you have to be able to get the GAL4 and UAS transgenes inserted into one of the chromosomes. In Drosophila this is often done by using a crippled transposon. A sequence of interest, for example the GFP coding sequence, and regulatory sequences to control its expression are cloned into a plasmid between transposon ends, most often the transposon is a P element. Along with your gene of interest, a marker is present between the transposon ends, often a gene that turns white eyes red (note eye color of the transgenic flies you will use in this experiment). This P element is injected into early embryos along with a helper plasmid that encodes the transposase. The transposase is transcribed and translated in the embryo and then recognizes the transposon ends in the other plasmid and inserts them into the genome. The plasmid encoding the transposase is diluted out and lost as the embryo develops, but if the crippled transposon has been inserted into a chromosome in a germ cell, then you can generate a line of transgenic flies. If you are interested in learning more about how this is done, one cool resource is: http://www.hhmi.org/biointeractive/clocks/vlab.html RNAi, and adaptations for use in Drosophila RNAi is a relatively new method to reduce levels of a particular protein in an animal or cell of interest. Basically recognition of a double-strand RNA by the cell triggers a mechanism that targets the endogenous RNA with the same sequence as the double-strand RNA to be degraded. It is pretty cool, and if you do not recall from previous classes how it works, then I suggest going back to your textbook, or looking at wikipedia (http://en.wikipedia.org/wiki/RNAi). It is a method that has been absolutely revolutionary! In practice, how RNAi is performed varies from species to species largely depending on which parts of the RNAi machinery are expressed in somatic cells of that species. For example, many mammalian cannot process long double-strand RNAs into the short 21-mers that trigger RNAi, so people use short hairpin RNAs that do not need processing. In Drosophila long dsRNAs can be efficiently process to the short fragments that trigger RNAi, so typically long hairpins are made. In neurons for these to be efficiently processed we do need to supplement the endogenous RNAi machinery by adding extra Dicer2, but other Drosophila cells seem to express enough that this needs to be added only in neurons. Large double-strand RNAs are made is specific cells in Drosophila using the GAL4-UAS system. In this version of it an inverted repeat is inserted downstream of the UAS sequence, so that when it is transcribed in response to GAL4 a hairpin RNA is made. In recognition of the power of this

From: An Introduction to Genetic Analysis, by Griffiths et al. In this case the transposon to be inserted in the genome contains the rosy eye color marker. The helper plasmid is the one that encodes the transposase.

approach several different consortia across the globe decided to make libraries of Drosophila lines that contained UAS-hairpin transgenes. We will use RNAi lines from one of these, the Vienna Drosophila RNAi Center (VDRC). This is a fabulous library and contains line that target almost 90% of Drosophila genes. Please see http://www.vdrc.at/rnai-library for a brief description.

Diagram of how RNAi is most often performed in Drosophila. Picture is from the VDRC web site:http://www.vdrc.at/rnai-library Other web sites you will need to make use of for this project: pubmed: http://www.ncbi.nlm.nih.gov/sites/entrez use this site to find papers that describe what the gene you are targeting does flybase: http://flybase.org/ this site gives a great overview of what is known about each Drosophila gene, and also has other helpful resources Background on dendrite shape: Dendrite development: http://www.ncbi.nlm.nih.gov/pubmed/19270170 Dendrite morphology and relationship to brain disorders: http://www.ncbi.nlm.nih.gov/pubmed/22465229

Extra protocols Live Imaging of Drosophila Larvae 1. Collect and age larvae (for RNAi experiments we typically put flies in a bottle with small holes poked in it, melt food into a 35mm cap, use cap to stopper the bottle and let flies lay embryos on it overnight. Then we remove the cap to a petri dish and age three days at 25 degrees). 2. Remove larva gently from food with old, fairly dull, forceps. 3. Put larva into a 35m dish filled with Schneiders Insect media (or water). 4. Allow food to wash off larva. 5. Remove larva from Schneiders gently with forceps. 6. Place in center of dried agarose pad on a slide. 7. Allow larva to start to crawl and become dorsal side up, then remove excess media with a kimwipe, but do not dry out completely!!. 8. Put a small piece of tape on one side of a 22X40 mm coverslip, press onto slide. Lower coverslip until it is just above larva. When larva is fully extended, press coverslip gently down (dont pop it!), tape the other side of the coverslip to the slide. This part is tricky, but super important. If the larva dries too much and gets stuck, add a little Schneiders to free it then start over. 9. You are ready to image! The larvae are pretty healthy using this method, but still try to look at them pretty quickly. If you want to save the larva and look at it again later, remove the coverslip and add a little Schneiders to free the larva. Then move the larva back into food. Materials needed: Box of glass microscope slides 3% agarose container to hold slides (while laying flat to dry) transfer pipettes 2 slides with tape in the middle of each slide (see picture)

Procedure: 1. Set up slides as seen below, with the slide receiving the agarose slide in the middle.

2. Heat 3% agarose to boiling (remove bottle with gloves) 3. Place a drop of liquid agarose onto clean slide with transfer pipette. 4. Use an additional clean slide to lightly press on the drop to even out the pad. (see below for set up)

5. Let the drop sit for ~3-5 seconds then slowly move slides apart, so that you end up with a flat drop on one of the slides, you can reuse the other one. 6. Place slide, agarose side up, in container. 7. Upon completion of all slides, place container in the oven (~65C) overnight to ensure pads are dry and cemented to slide (or just leave on bench- this should work too!).