Microinjecting worms

互联网2013-12-26

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<center> <h2> Microinjecting worms </h2> </center>

<center> by Michael Koelle 8/23/94 </center>

You will have a couple frustrating sessions when you first attempt this technique, but everyone seems to master injection after a few days, and it works very quickly and reliably once you have some experience.

1. Materials
a) agarose pads : make a lot of these at a time; they last forever.
Using a Pasteur pipette place a drop of 2% agarose in H20 on a 24x50 mm coverslip. Drop a second coverslip on top, which will flatten the agarose into a thin pad. (try to avoid air bubbles, but a few won't hurt anything) When the agarose has hardened (> 5 sec) slide off the top coverslip. Use this top coverslip as the bottom coverslip to make the next pad; its thin coating of agarose will make the pad stick to it instead of the fresh top coverslip.
Place the coverslips in a box and cover with aluminum foil to dry. Can leave out on the bench overnight, or bake in an oven at 65・ (the same one you use to de-mite worm boxes) for 1 hour, or bake in an 80・ oven for about 15 min. Once dried, you can store the pads by sticking them back in the original coverslip box.
b) 10X microinjection buffer .

20 % polyethylene glycol, molecular weight 6000-8000
200 mM potassium phosphate, pH 7.5
30 mM potassium citrate, pH 7.5

Mix 10 mL 1M K phosphate pH 7.5, 5 ml 300 mM K citrate, pH 7.5, 10 g PEG, and ~25 ml H20, stir ~10 min to dissolve the PEG, and add more H20 to final volume of 50 ml. Note that the PEG is very near its solubility limit, so the solution may remain cloudy until the solution is vol'd to 50 ml with H20.

Making the buffers: 1 M K phosphate pH 7.5

8.7 g K2HPO4 + 50 ml H20 = 1 M solution
6.8 g KH2PO4 + 50 ml H20 = 1 M solution
mix 32.4 ml 1M K2HPO4 + 7.6 ml 1 M KH2PO4 to get pH7.5

300 mM K citrate, pH 7.5
6.3 g citric acid + ~70 ml H20
add HCL or 10N KOH to pH to 7.5
H20 to 100 ml

c) recovery buffer : M9 buffer. People used to use M9 plus 4% glucose; the glucose is unnecessary and only causes the solution to become contaminated.
d) Microinjection needles : use a microscope slide box to store pulled needles. Put two strips of modeling clay in the box to hold the needles (press them into the clay, leaving the tip hanging free in space.
We use "Glass 1BBL w/FIL 1.0 mm 4 IN" filaments, item #1B100F-4 from World Precision Instruments, Inc., (813) 371-1003, FAX (813) 377-5428. Keep these clean, always immediately recap the tube after removing a filament.
We use a Kopf needle/pipette puller Model 750, from David Kopf Instruments, Tujunga, CA. Turn the machine on (switch at back right). Push the button in the back to reset the programs and get "0000" displayed. Flick the lever to "program", press "b", then press the program number being used then "e" for enter. Pressing "e" successively will tell you the parameters set by the program. Flick the lever to "run".
On the Stern lab machine we're using program 10, which is: heat1=5 AU, heat2=0 AU, sol= 5 A, delay= 0 sec, sol= 0.1 sec. It takes 7 to 15 seconds to pull the needle, although the best needles are usually pulled in 11-12 seconds. It is our practice to not allow each individual to adjust the spacing of the filaments (which is done using the middle set of knobs). Individuals can adjust the machine to their preference by using different programs. This allows everyone to reproducibly pull needles they like without a fuss.
Insert a glass filament into the needle puller without touching your fingers to the part that will be heated, or touching the filament to the heating elements. Tighten the filament in with the top knob, DON'T EVER TOUCH THE MIDDLE KNOB!!!! Slide the bottom unit up all the way, then tighten the bottom knob so that the unit is held suspended. The green "ready" light should now be on. Close the cover.
Press the "start" button. The machine will time how long it took to pull the needle; want it to be ~10.4 sec. Carefully remove the bottom half of the filament in the box with clay, discard the top half of the filament. I pull about 8 needles at once. Lately, we've been putting the filament in higher in the machine and taking the top needle.
e) Microinjection oil : we use "halocarbon oil series HC-700", P.O. number 030B31793, 1 lb bottle, from Halocarbon Products Corporation, 130 Dittman Ct., N. Augusta, S.C. 29841. 9/93: the bottle now says CAS #9002-83-9. The Hlocarbon Products Corporation address is P.O. box 661, River Edge, NJ 07661. Phone number: 803-278-3500.

2. Making the DNA solution
Want clean DNA buffered at pH 7.4 in a K+ buffer, with not too much Na+ in it. Up to 25-40% DNA prepared using Qiagen columns in TE, made up in 1X injection buffer (see above) is okay. If necessary to get rid of Na+, can make the DNA 0.1 M KAc pH 7.4, add 2 vol. EtOH, ppt, wash in 70% EtOH, and resuspend in 1X injection buffer. (In one experiment, I injected a 40% TE mix and got 20 F1 rollers from 30 injected animals. Then I ppted the DNA and resuspended in injection buffer, injected again, and got 70 rollers from 15 injected animals. With other DNAs I've also noted several-fold better results using DNA in injection buffer than I have typically gotten using DNA in TE. It therefore seems very worthwhile to use DNA in injection buffer.)
Typical DNA concentrations: When trying to rescue a mutant with cosmid pools, use pRF4 (contains the dominant rol-6 mutant) at 80 µg/ml, and each cosmid to be coinjected at 20 µg/ml. Some cosmids contain poison sequences; in this case no transmitting F1s will be generated. Michael Stern had this problem with sem-5 and solved it by reducing the cosmid concentration to 1 µg/ml. For ß-gal constructs, coinject pRF4 and the plasmid construct each at 80 µg/ml. Some constructs exhibit dominant phenotypic effects. This problem has been solved in some cases by lowering the concentration of the ß-gal DNA construct injected.

3. Setting up the scope, loading the needle, mounting, and breaking the needle.
Set up the scope and inector: We use a Zeiss Axiovert 10 microscope; the relevant objectives are the plan neofluor 5X and 40X. Mounted on the scope is a Narishige Model MO-202 micromanipulator, on which is mounted a needle holder hooked up to a Narishige IM300 injector. A nitrogen gas tank is hooked up the microinjector.
To inject, turn on the N2 gas tank. Use the regulator to adjust the pressure coming from the tank to about 75-80 psi; it should already be set to this range and should not require Turn on the microinjector. After a few seconds the display will show the pressure the injector is receiving from the nitrogen tank. It should be about 75-80 psi (if not, adjust the regulator on the N2 tank to get it in this range. Next adjust the pressures used for injection: press the"mode" button twice until the display shows the four pressure settings (fill, inject, balance, hold). We don't use the fill or hold settings - ignore these. Using the silver knobs on the injector, adjust "inject" to 18.9 psi to start with. Adjust the balance pressure to about 2.5 psi. During injections, you will switch between the balance pressure and the injection pressure using the foot pedal. The balance pressure, used between injections, is a constant low pressure level used to keep the injection oil from backing up the injection needle by capillary action. The higher injection pressure is used to pump the injection solution into the worm. The injection and balance pressures can be adjusted to suit the needs of the particular needle you are using. For example, a needle with a very small opening may require a higher inection pressure to get an adequate flow of the injection solution. After adjusting the pressures, press the "mode" button three more times until "action" appears in the display. Press the "baln" button to turn on the balance pressure. You can now toggle between the balance and inject pressures by pressing the foot pedal. If the needle becomes clogged, you can try clearing it by pressing the "clr" button, which is currently programmed to give a 1 second pulse of high pressure (the same pressure coming from the N2 tank, ~80 psi).
Loading the needle: Before loading the needle, microfuge the DNA solution for 10 (some say 30) min, to pellet particulate matter that might clog the needle. Place a 0.5 µl drop of the solution on the back (unpulled) end of the needle; the needle contains an inner glass filament that will wick the DNA solution to the other end. Holding the needle up to the light, you should see liquid at the tip of the needle. Look at the needle under the dissecting scope to see if there are any air bubbles trapped in the liquid. These are bad; for some reason blowing them out through the tip often causes the needle to block, perhaps dirt adheres to and maybe causes the formation of the bubble in the first place. If the bubble is truly tiny and near the needle tip, proceed to break the needle and blow the bubble out the tip. If the bubble is bigger, mount the needle on the scope, tilt the needle so the tip is pointing as nearly straight down as possible, and go away for ~10 or more minutes. Hopefully, the bubble will rise up out of the needle tip into the liquid resevoir above the taper of the needle where it is harmless.
Mount the needle on the scope: The whole top part of the axiovert tilts back so that you can get at the needle, and also so that you can change slides on the stage without risking touching the needle. Remove the old needle by unscrewing the assembly that holds the needle. Be very careful here ; the pressure can cause the needle to shoot out like an arrow here, so keep your face etc. out of the way. Also, there are two small black rubber O rings in the assembly; make sure these don't fall out and get lost. Remove the old needle and throw it out (you will leave your needle in when you're done) and insert your needle (back end first so as not the break the tip, obviously), and tighten the needle by screwing the assembly together well. Leave a few millimeters of the back end of the needle sticking out the back end of the assembly. Then screw the assembly on to the holder on the scope (don't do this as tightly as you screwed together the assembly itself- that way when you take the needle off next time the whole assembly will come off as a unit and the needle won't shoot out like an arrow.) Turn the three knobs on the fine control of the micromanipulator to the middle of their range (5), and using the course controls (knobs on the part of the micromanipulator mounted on the scope), make sure the needle is high enough so that when you lower the top half of the axiovert, the needle tip won't crash into the stage. Also use the coarse controls to move the needle tip left/right forward/backward until it is just above the objective (will then see it glowing in the light shining down from the condenser).
Breaking the needle: There are two methods to break off the tip of the needle. The (older?) method of etching the needle tip with hydrofluoric acid is falling into disuse in the Horvitz lab, and the acid is somewhat dangerous.
The more common method is to physically break the needle. Over a Bunsen burner draw out a standard (not microinjection) 10 µl micropipette to about 1/5 its starting thickness. Place a stretch of the drawn out part on a 24x50 mm coverslip, and put a drop or two of microinjection oil on top. Mount this on the axiovert, and using the 5X objective, focus on the micropipette (see a sharp black line on the edge when you are focused on the middle). Jin suggests putting the micropipette so that it doesn't go straight up and down, but rather is at a slight (30・?) angle, so as to get a beveled edge on the microinjection needle. Using the fine controls, carefully lower the injection needle towards the stage until it is in the same focal plane as the micropipette. At this low power, you can't see the actual tip, so you may have to try the 40X objective to do this. Again, using the fine controls, slowly move the injection needle left until it touches the micropipette, and then pull it back. To check the needle, press the foot pedal to look for flow out of the needle. Should see pretty rapid flow out of the needle using pressure P2, but none at pressure P1. If there is no flow at P2, the needle isn't broken; try again. You will have to see by experience what the optimal flow rate is. You want to be able to flood the gonad in about 3 seconds of flow at P2. You can make fine adjustments to the flow by adjusting P2 with the knob. When done breaking the needle, use the fine control to lift the needle up out of the oil in preparation for injecting.

4. Mounting worms on an injection pad.
Take out an agarose pad and breathe on it (about 1 long breath) to moisten it; if it is too dry the worms will dry out and die - too wet and the worms won't stick well. Place a drop of microinjection oil on the pad. Lay the cover slip on the top of an upside down lid of a small worm plate with two strips of lab tape across it. This holds the cover slip at about the same height as the worms on a plate so that you don't have to focus around too much when switching back and forth. I like to spread the oil drop around with a worm pick so that the oil isn't too deep.
Using a worm pick with oil on it as glue (want to minimize the amount of bacteria you transfer) transfer adult hermaphrodites to the oil drop on the pad. If there is still adhering bacteria, push the worms around in the oil with a pick until the bacteria come off. Most people like to use first day adults that have a line of about 10 eggs in them; these have large robust gonads. Jin picks L4 animals and ages them one day at 20・ before injecting. For Egl animals, you have to inject them younger before they become bloated.
As a beginner, stick 1-2 animals on a pad. Some experts do up to 9 animals at once; I prefer to do only 2-3 to minimize the time (and therefore trauma) that the worms spend drying out on the pad during injection. The trick is to stick the animals down in the correct orientation so that the vulva is pointing to the side, and the two distal gonad arms (the syncytial part which you will inject) are up against the wall of the animal on the opposite side from the vulva. You don't want the syncytial gonad to be on top or underneath the animal. When the animal is in the oil, the syncytial gonad is visible as two clear areas towards the anterior and posterior of the animal. To stick the animal right, wait until it is floating in the oil so that it's body flexures go sideways, not up and down, and pat the animal down on the agarose pad with your pick until it is stuck to the pad. Avoid stroking or patting the animal on the head, which can kill it; ideally the animal will be fully immobilized except for it's head, which will still be free and wiggling. The animals stick best when they first touch the pad; if you fail to stick them on the first try, it becomes increasingly difficult to stick them down; after you've rubbed them all over the pad apparently the stuff that allows them to stick to the agarose becomes worn off. Sophisticates can stick down a whole set of animals in a line in the same orientation for assembly line injecting. Once the animals are in the oil, work reasonably fast to get the procedure over with before the animals dehydrate.

5. Injecting
Rotate the top of the microscope back, place the coverslip on the stage (don't use clips to hold it on, you can remove the clips from the stage). Carefully lower the top of the microscope, watching the needle to see that it doesn't crash on the coverslip (if you raised it a bit off the stage before, this won't be a problem). Alternatively, I like to just raise the needle fairly high with the micromanipulator in between injectiions, and then slide the old coverslip out from underneath it, and slide the new one in. Using the 5X objective find the worm, make sure it is in the correct orientation (vulva away from the needle). Can move or rotate the entire stage to move the worm, although some like to move the coverslip itself. It is best to have the worm at a 45・ angle to the needle; this maximizes the path length for the needle inside the gonad, helping to make sure you get the tip in the gonad instead of going all the way through and out the other side. Carefully lower the needle into the focal plane with the fine adjuster (at this point, you only need to move the needle up and down with the micromanipulator; you always move the worm, not the needle, up/down left/right, by moving the whole stage).

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