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Koelle Lab Protocol_microinjecting worms.html


Microinjecting worms

by Michael Koelle

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).
Move to the 40X objective. Focus on a syncytial gonad arm; this is recognized as a sausage shaped clear area surrounded by nice round nuclei. Kimble and Sharrock (Dev. Biol. 96:189-196 (1983)) show an excellent photograph of a dissected gonad that should give you a good idea of what to look for if you're new to worm anatomy. Focus on the center of the sausage so that you see a nice row of nuclei on either side of the sausage. Using the fine adjuster, move the needle up/down until its very tip is in focus. Gently move the worm so that it is pressing gently against the needle at a point where the syncytial gonad is pressed up against the body wall, and so that the needle tip will end up inside the gonad after it penetrates the body wall. To penetrate the body wall, use your right index finger to gently tap the micromanipulator on the little box with the ball joint in it (just above where the arm the needle is on is attached). This vibrates the needle a little so that it punctures the worm. Hopefully the tip is in the gonad now; if it obviously isn't pull out and try again.
Press the pedal to start the flow of DNA. If you're in the gonad it should be obvious; as the gonad is flooded it bloats like you're filling a sausage, and you can sometimes see the nuclei in the syncytium reacting to the flow. You want to put as much liquid in the gonad as possible; hopefully it will flow all the way around turn of the gonad. Eventually the gonad gets so huge that liquid starts to blow out the animal through the hole that the needle went in; try to avoid this but it's ok if this happens - you want to load the animal about to this point. A good rule of thumb is to inject until you see a good amount of liquid has made the turn and has flowed into the proximal gonad, and then to shut off the flow. To stop the flow press the pedal again and move the animal away to get the needle out. Mello et al. (EMBO J. 10: 3959-3970, 1991) show excellent photographs of a gonadal flood.
Usually one gonad arm is much easier to see well than the other, so some people only inject the easy gonad arm. Others try to inject both. If you miss the gonad, you will see liquid filling the pseudocoelom. Usually, the animal is ok, and you can just try again. It is surprisingly hard to kill the worm by jabbing and injecting it incorrectly.
Some people (Jin) press the P1 button after every injection to clean the needle and help keep it from clogging. Eventually needles tend to clog and must be changed. After you finish a worm, use the fine controls to lift the needle out of the oil before moving the stage to find a new worm, or removing the pad.

6. Recovery
Put the pad under the dissecting scope (on the inverted plate lid) and using a P200 pipetteman place a drop of recovery buffer on the oil drop above the worm. Then poke a worm pick straight down through the recovery buffer and oil to touch the agarose pad next to the worm. This will form a channel, and the recovery buffer will form a layer underneath the oil in which the worm will float. Worms can be left on the pad in recovery buffer for hours, but you might as well immediately move them to plates. (Some say it is better to leave them in recovery buffer for > 5 minutes - in this case place the coverslip in the lid of a large worm plate, and place the plate over it to make a humidified chamber.) Can put up to 3 injected worms on a plate; I prefer one worm/plate. Use a slightly drawn out and broken off and flamed smooth large diameter micropipette (1.5 mm diameter drawn out to about half that) and mouth pipetted the worms over to a plate, and set a 20·.

7. Results
Three days after injection, score the F1 for the marker gene phenotype (e.g. rollers if pRF4 (rol-6) is used). rol-6 animals are Rol even as young larvae, so it is tempting to score and pick the F1 after only two days: don't do this! The young larvae are very delicate and you are liable to kill them by picking them. Each Rol F1 is considered an independent transformant (even if several come from the same injected P0). Therefore, each Rol F1 should be placed on a separate plate to try to get lines.
Typically people inject 30 P0s (takes just 2 hours if you're good), and expect to get 3-300 Rol F1. Usually some of the injected P0s give zero or 1 Rol F1, most of the P0s give 5-15 rollers. As a beginner I averaged 1-2 F1 rollers per P0. Now I average about 8 per P0, and some people do much better. Of the Rol F1, typically about 5-30% will transmit the array, allowing a line to be established. Typically, lines transmit the array to 30-80% of their progeny. There is variation among lines transformed with the same DNA. For example, only a fraction of lines transformed with a cosmid/pRF4 might give rescue of a mutation in a gene found within that cosmid, and the strength of the rescue will vary among lines that do show rescue. In the Horvitz lab, people look at ~6 lines before they tentatively believe a negative result.
Some lines transmit at only a few percent per generation. The frequency of transmission varies from animal to animal. Jin says to be sure to keep these lines when injecting ß-gal or GFP constructs; the low transmission rate of the extrachromosomal array is useful when trying to select for chromosomal integration of the array (leading to 100% transmission).
Some people (me, Mark , Gillian, Jin, Tory) have noted a high incidence of males in the F1 of injected animals, or in some Rol lines. Since this was observed using a variety of different DNAs it is likely a nonspecific effect of extrachromosomal DNA on chromosome disjunction, and doesn't mean your gene is involved in sex determination.
If you are rescuing a mutant, and using pRF4 as a coinjection marker, you may notice an odd effect; a high proportion (up to half) of non-Rol F1 progeny of rescued Rol animals may themselves also be rescued for the mutant phenotype. This is not necessarily indicative of maternal effect rescue of your mutant. Rather, it can be due to lack of penetrance of the rol-6 dominant allele and/or mosaicism for the extrachromasomal array. This can be demonstrated by picking individual non-Rol rescued animals and showing that they throw Rol progeny.
8. Alternate coinjection markers
Sometimes it is not desirable to have the dominant Rol phenotype in your transgenic worms. In these cases, you can microinject worms carrying a recessive marker mutation with a rescuing plasmid for that gene, along with whatever your experimental DNA is. The following properties are desirable for such a marker mutant: 1) it should be very easy to score, preferably at all stages of development. 2) the mutants should have healthy gonads that are easy to inject. This is sometimes achieved by using a ts allele, growing the animals at the permissive temperature before injection, and then shifting to the nonpermissive temperature. 3) it should be possible to get strong F1 rescue of the mutant. 4) the rescued animals should be truly wild type.

I've heard about people using unc-76, dpy-20, and lin-15 as coinjection markers. I've been using lin-15. Its main drawback is that it can only be scored in adults. lin-15(n765ts) animals are raised at 15· for injection. A lin-15 rescuing plasmid is included at 50 ng/µl in the injection mix. I'm using the plasmid pL15EK, which I got from Xiaowei Lu. This is an 11 kb Eag1/Nru1 rescuing fragment of cosmid C29B12 cloned into pBSKS+ cut with Eag1/Kpn1, (using a Kpn1 linker on the Nru1 end). After injection, the worms are moved to 20· or 25·. At 25· the non-rescued animals are very sick, and there is a strong selection for transgenic worms. At 20· the worms are healthier, and the transgenic worms are recognized as non-Muv. You should wait 4 days after injection at 20· to score the adult F1. Even though the Muv phenotype only develops during the L4, it appears that at 25·, n765 animals reach the L4 stage more slowly maternal rescue. People usually put lin-15(n765)/+ animals at 22.5· in order to enhance the Muv phenotype of the n765 homozygotes they throw. In two trials using lin-15 as a coinjection marker, I got about the same number of F1 non-Muv animals as I usually get F1 rollers using rol-6 (i.e. about 100 F1's from 20 injected P 0 s). However only 4% and 8% of these F1s transmitted their array, whereas typically more than 20% of my Rol F1s transmit. Piali in the Bargmann lab says she gets about 15% transmission; she's using a different lin-15 plasmid than I am.
I find that lin-15 is much preferable to rol-6 as a marker when trying to integrate an array. The Muv phenotype is incredibly easy to spot, whereas screening plates for the absence of non-Rol animals (which you do when trying to integrate pRF4) is a lot harder.