Bier Lab DNA Injection protocol：比尔实验室DNA注射协议

Bier Lab DNA Injection protocol：比尔实验室DNA注射 协议 离婚协议模板下载合伙人协议 下载渠道分销协议免费下载敬业协议下载授课协议下载 Bier Lab DNA Injection protocol Heather Elledge February 2007. Materials and Reagents , Qiagen Qiafilter Midi Kit (Cat #12243) , 10X Injection Buffer (IB) , W- or w1118 flies, young , Two small fly cages , Small grape plates , Yeast (fresh, no more than two weeks old) , Glass slides with double-sided sticky tape (Scotch 3M brand) , Large Petri dish with a bar of clay running through the middle , Glass capillaries (A-M Systems Cat #615000) , Desiccant (Drierite Cat. #23005) , Halocarbon 700 Oil , Empty plastic slide box (Fisher Cat. #03-448-5) , Pick/Forceps/Pencil with a sewing needle in the eraser , Fresh fly food in vials A. DNA Preparation: 1. Purify the DNA/vector using the Qiagen kit. Follow the instructions to completion (includes an isopropanol and 70% ethanol wash) and record the concentration using a spectrophotometer (OD). Do this for p,2-3 too, 260 which is needed to successfully transpose the P-element into the germ-line. 2. Do not perform a phenol cleanse as this is toxic and may harm the embryos. 3. In a separate tube, add ,2-3 and DNA in a 2,g:8,g ratio (roughly a 1:5 ratio of ,2-3:DNA) to where the final volume (in water) is 9,l. To this add 1,l of 10X IB. 4. Spin for 10 minutes at 14000 rpm at room temperature. 5. Transfer 8,l of final product to a new tube, being careful not to touch the bottom of the original tube. This new tube now contains pure construct and can be injected. 10X Injection Buffer Recipe: 50ml -100 mM Tris, pH 7.5 5 ml 1M, pH 7.5 -1 mM EDTA 1 ml 0.5M, pH 8.0 -100 mM NaCl 10 ml 5M -Use ddHO, does not have to be filtered add DDW to 50ml 2 -Autoclave twice, 20min each with cooling in between -Store at 4?C -pH should be about 7-8, food coloring is optional B. Cage Preparation 1. In order to make successful cages of flies that will lay all day, the w- stock must be amplified in bottles or vials. Two to three days before scheduled injections, make two small cages with flies that are less than five days old. Small cages can be found in the bottom drawer of the bench containing the Denville centrifuge. Stuff the large hole in the side with a cotton ball (this allows the flies to breathe). 2. Prepare fresh yeast in a 50ml centrifuge tube by mixing Active Dry yeast (Genessee cat #62-103) with dHO from the sink (foot pedal). The yeast 2 should have the consistency of chunky peanut butter—anything too creamy will cause the flies to stick and drown. 3. Grab previously prepared (by the dishwasher) grape plates from the cold room and spread a thick amount of yeast product in the middle. In the days leading to injections the older grape plates (over a month) can be used, but on the day of injections be sure to use the freshest plates. Also, in the days leading to injections there should be a good amount of yeast on the plate to keep the flies happy, but during injections there does not need to be a lot. 4. Keep cages in the dark either in room temp or in the 25?C incubator, making sure the grape plates are changed every morning and evening. 5. On the morning of injections, transfer all flies to new cages. Do not anesthetize them with CO. Transfer grape plates every half hour. 2 C. Prepare slides with double-sided sticky tape more than 24 hours before injections, as tape fumes are toxic. Sticky-tape D. Needle preparation 1. Place a single layer of paper towel on either side of the clay bar in the Petri dish and saturate with water. This will keep the chamber humidified. 2. Pull about 10 needles using the puller in the microinjection room (BH4325), making sure only two of the four weights are attached. Carefully remove the lower needle and place in the Petri dish, sticking it to the clay. Discard the top needle. Be sure needle tips point in same direction. 3. With a P2, P10, or P20 pipette, add ~1,l of DNA to the top of the needle, allowing a bubble to form which will break as the fluid is drawn to the tip. 4. Once all fluid has collected at the tip, attach a needle to the tubing and place in the holder of the microscope. Adjust set-up to bring the tip into focus, then raise needle and place a slide onto the base, bringing it and the needle into the same focal plane. The slide edge should be focused toward the center of its thickness, not its top or bottom. This will cause the slide edge to look blurry. Slide Edge of slide Tip of needle 5. Attach the other end of the tubing to the 20CC syringe and press on the syringe to force DNA toward the tip. SLOWLY bring the slide edge to the needle until contact. Let a small drop of DNA exit, then quickly pull the slide away. The needle tip should be sharp and narrow. If needle is too blunt toss it, because it will slaughter the embryos. DNA Broken needle tip 6. Leave needle tip in halocarbon oil between each injection slide to keep DNA at the tip from drying. Release syringe from the tubing between each injection to keep the oil from getting drawn into the needle. E. Other Preparations Necessary 1. Change the Drierite desiccant in the Petri dishes the morning of injections. The pellets should be blue. If purple, pink, or white, it is too old. 2. Prepare incubation chamber by placing paper towels in an empty slide box and saturating with water. 3. Check the 50ml centrifuge tube in the microinjection room. Make sure there is about 5ml of halocarbon oil in it, and place the glass stirring rod into the tube. 4. Don’t forget your timer. F. Embryo Collection and Rolling 1. Upon entering the lab, change the grape plates on the cages. Wait 30 minutes and change the grape plates again. Throw out the first set, as the embryos are too old. Wait another 30 minutes to change plates. This time, the grape plates from the cages can be used for embryo collection. 2. Use a pick/forceps/sewing needle to carefully pick embryos off the grape plates and place them on the left side of the tape. 3. Gently push/massage the side of each embryo to pop it out of its chorion. Pushing from up top could smash the embryo. Once out of its shell, roll it back over the chorion, off the tape, and the embryo will stick to the pick. 4. If the embryo is transparent in any way, kill it for it’s too old. If the embryo is bent in any way while trying to pop out of the chorion, it is dead. 5. Align the rolled embryos in the middle of the slide, making sure all of the anteriors are facing the same direction. Keep them an embryo width apart. Needle Direction 6. Each slide should only take 15 minutes, to allow for two slides per grape plate change. 7. Place slide of rolled embryos into the Petri dish full of desiccant, making sure the top of the slide does not connect with the lid (embryos will get squished). Desiccate embryos for 10-11 minutes, depending on the level of humidity in the injection room. G. Injecting 1. Once the timer beeps, take the slide out of the desiccant. Be sure to keep the lid on the desiccant dish, as oxygen will turn the pellets purple and render them ineffective. With the glass stirring rod, cover the embryos with a small layer of halocarbon oil, taking care not to let a drop cross the tape onto the slide. Good Bad 2. Raise the needle and exchange the slide used to break the needle for the slide with embryos. Bring the posterior of the embryos into focus, and then attach the 20CC syringe to the tubing. Carefully lower the needle into the oil until the tip is in the same focal plane as the posterior of the embryos. Anterior of embryo Needle Micropile Posterior of embryo 3. Gently squeeze the syringe to check DNA flow. The tip should not be blunt, but still allow flow with barely any pressure on the syringe. Optimally, the syringe could be let go, and the DNA could still flow. 4. Slowly poke the posterior of the embryo with the needle. Take care not to allow the needle to penetrate too deeply into the embryo (beyond a third of the embryo’s length). As soon as you see the DNA enter the embryo, pull out. Too much DNA can be toxic. Optimally, all DNA should be in the most- posterior portion of the embryo. If the embryo explodes, spilling cells upon contact, the slide was not desiccated long enough, increase the time. If the needle pushes the embryo into itself beyond a third of its length before penetration, the slide was desiccated too long, reduce the time. 5. Some cells can exit from the hole made by the needle. Use the needle tip to draw the bubble away from the embryo. The halocarbon oil will act as a plug and prevent more cells from escaping. This will save the embryo. 6. The embryo should look brown. If any part of the embryo is darker, destroy it, for it has already died due to bending while rolled. If the edge of the embryo is transparent or any cells have differentiated, kill it for it is too old. If the embryo is pointed in the wrong direction, its micropile facing the needle, destroy it. Injecting DNA into the anterior is toxic and will kill it. 7. Taking these situations into effect, it is possible to have to destroy half the embryos on the slide. Don’t worry. If over 12 slides are rolled and injected, there will be enough embryos. It is better to kill any embryos that have no chance at a successful transformation than allow the researcher to waste his/her time on a cross that was doomed from the start. 8. Continue injecting DNA into the rest of the viable embryos on the slide. 9. Once finished with a slide, leave it on the microscope with the needle still immersed in the oil. Release the 20CC syringe from the tubing. Do not place the slide into the incubation chamber until a new slide is ready to be injected. H. Incubation and Larval Collection 1. When injections are complete, all slides should be in the incubation chamber. Turn off microscope, take stirring rod out of halocarbon oil and close the 50ml centrifuge tube containing the oil, and discard all needles into the red sharps container. 2. Allow the embryos to incubate in the microinjection room overnight before taking them into room temperature for larval collection. 3. Use fresh fly food, prepared that day, for all larval collection. Cut a cylinder out of the food or mash up the top with a spatula. Using a dissecting microscope and pick, look for any larvae swimming in the oil. If it is apparent that an embryo is alive, but is still attached to the sticky tape, do not remove it yet, for it can easily be destroyed. Scoop up the floating larvae using the pick and gently place them onto the fly food. The larvae could be squished at this step as well, so use caution. 4. Leave the chamber at room temperature, checking once or twice daily until only dead embryos remain. This could take up to three days, so plan the injection date accordingly. Tuesday or Wednesday is the best day to inject. 5. Place all vials containing larvae into the 25?C incubator, or leave at room temperature (either is acceptable). Start collecting male and virgin female w- flies for crosses. 6. At room temp it will take about 14 days for the flies to eclose, while at 25?C it will take 8-10 days. Cross each female individually with 2 w- males, and each male individually with 5 w- virgin females. Once the progeny of these crosses have eclosed, look for any with red eyes. 7. If found, jump up and down and cheer, for the injection was a success!