Sos Protocol
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Sos Protocol
Antisera staining/Immunohistochemistry
Fixation
AMPA assay
C. elegans cosmid vectors
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Sos (volatile repellent assay, smell on a stick) Originally developed by Cori Bargmann and published in Troemel, et al... Here you will find our version - this is very, very detailed. It should work for other volatile repellent chemicals, but we haven't used them yet.
Shinya Matsumoto (1) and Anne Hart(2)
(1) Research Institute for Food Science, Kyoto University, Gokasho Uji Kyoto Japan 611-0011, (2) MGH Cancer Center
149-7202 13th Street, Charlestown, MA 02129
an eyebrow hair attached to glass pasteur pipette (narrow end) with tape
NGM/agar plates which have been left at room temperature for 1-2 week (Tip1)
N2 (wild type) worms as positive controls (Tip2)
eat-4 (ky5) worms as negative controls
glass capillary pipette (optional)
1. Prepare 10 testing plates and number them on their lids. (#1, 10 for N2; #2, 9 for eat-4 ; # 3-8 for test worms)
2. Take 20-40 ul octanol to a microfuge tube. Dip an eyebrow hair 10-15 times into octanol to initialize the hair with the solvent. (Tip4)
3. Transfer 5-8 N2 worms to #1 testing plates by a pick. Try to pick worms on the outside area of the bacteria lawn and try not to transfer bacteria to the testing plates. Immediately after the worms were transferred to the testing plate, start the timer and write down the starting time on the assay sheet (starting time; 0:00) (Tip5)
Option: to transfer worms, you can try mouth-pipetting with SM buffer and a glass capillary pipette instead of using a pick. Sometimes transferring worms by a pick can damage the worms and it drive the worms to
strange behavior (for example, N2 not responding to octanol). Fill the glass capillary pipette with SM buffer and suck the worms into the capillary pipette by gently blowing out and into the solution. In this case to, avoid sucking bacteria.
4. Transfer eat-4 worms to #2 testing plate as in step 3. Write down the transferred time on the assay sheet after the worms were transferred.
5. Transfer testing worm to #3-8 plates as in step 3. Write down the transferred time on the assay sheet after the worms were transferred.
6. Transfer eat-4 worms to #9 testing plate as step 3. Write down the transferred time on the assay sheet after the worms were transferred.
7. Transfer N2 worms to #10 testing plates as in step 3. Write down the transferred time on the assay sheet after the worms were transferred. (Tip6, Tip7)
8. At time 10:00 (you should give worms at least 10 minutes for them to get used to the new environment), bring #1 plates under the microscope. Take a look at the worms and find a worm moving forward to test. Take off the lid and keep watching the worm you selected. The worm usually moves backward instantly upon removal of the lid, but in 5-10 seconds, it will resume normal forward movement. If the worm keeps repeating forward and backward movement, or does not resume normal forward movement, select another worm to be tested or wait until the worm has relaxed and resumes normal forward movement. (Tip8, Tip9)
Dip the hair once into octanol. Under the microscope, bring the tip of the
hair as close as possible to the head of the worm without touching the worm. When the hair is close, start counting until the worm moves backward to avoid the octanol. Write down on the assay sheet how long the worm took to start backward movement.
Do not test multiple worms!! Test one worm per plate. Any worms that have been tested, except the very first one, might have been sensitized or desensitized upon exposure to octanol during stimulation of the first worm. (Tip11, Tip11, Tip12). Give them a chance to recover. Close the lid. (Tip4)
9. Test the rest of the plates (#2-10) as in step 8. Make sure to start stimulation 10 minutes after the worms were transferred to the testing plates.
10. After all the ten plates were tested, return to # 1 and repeat assaying. However, take at least 5 minutes between assaying the same plate.
11. Repeat step 10 as many times as possible to obtain data. However, worms may not respond to octanol 25-30 minutes after they
were transferred to the testing plates. So, do not test worms more than 25-30 minutes after they were transferred to the testing plates.
12. Calculate the average and SEM (standard error of mean) by the following formula:
SEM = S (standard deviation of the data) / square root of n (number of the worms tested) (Tip13).
Tip1. The dryness of the testing plates does not affect the Sos assay as much as it does the Osm assay, but too wet plates sometimes affect the behavior of the worms. If the plates are too wet, worms will not respond to octanol. Take a look at worms 10 minutes after they were transferred to the testing plates. If you see water along the edge of the body of the worms, the plate is too wet. You should avoid using such plates. Plates that have been left at room temperature for 1-2 weeks after they were made should be fine to be used.
Tip2. For N2 and eat-4 , the temperature that they have been raised at does not affect the behavior of the worms. However, N2 or eat-4 raised at 15 Celsius degree have not been tested.
Tip3. It does not matter whether the SM buffer used for mouth pipetting does or does not contain cholesterol.
Tip4. Cut off the tip of the hair that has been attached to a glass pipette
so that octanol is absorbed easily into the hair (or the hair is thicker and absorbs more). When dipping the hair into octanol, take care not to let the tape touch the octanol. Perhaps ingredients in the tape could be dissolved by octanol.
Tip5. If bacteria was transferred to testing plates along with the worms, they will tend to stay at the location of bacteria and will not respond to octanol.
Tip6. N2 and eat-4 are tested at both the beginning and the end of the cycle to confirm that worms are responding to octanol throughout the assay time.
Tip7. The original plates that worms are picked from should be returned to the incubator after all the required worms are transferred. I do not know why but I have sometimes had trouble with worms (including N2 and eat-4 ) that had been left on the bench for a while.
Tip8. I have occasionally found that N2 worms do not respond at all to octanol. In such case, it is useless to use other worms from the same plate in the assay because they are often also insensitive to octanol. If you find insensitive N2 worms, just throw the plate away. I have found that even if the parent N2 worms are insensitive, their progeny are sensitive and and can be used as controls. On the other hands, I never found that all eat-4 worms from a same plate becoming sensitive to octanol.
Tip9. If you can feel the air moving (or turbulence) around you (around the testing microscope), it could influence the behavior of the worms. They keep changing forward and backward movement rapidly and they take much longer (30-90 seconds) until they are settled and resume normal forward movement. If you experience this, move to another scope.
Tip10. Bring the hair as close as possible to the head of the worm. Do not be afraid of touching the worm with the hair. If you touch the worm, get rid of the worm immediately. You have to practice a little to be able to bring the hair very close without touching the worm, but you will get used to within short time.
Tip11. If you touch the surface of the agar plate with the hair during stimulation, the worm reverses probably due to the shock of the touching, and not due to avoiding octanol. Do not count these trials. In addition, it will result a drop of octanol formed on the agar surface. Keep the lid open for a such plate until next cycle (5-7 minutes after) for octanol to evaporate and for worms to be recovered from octanol-exposure.
Tip12. Start counting exposure time when worms are exposed to octanol and therefore can respond. This timing is somewhat objective and you have to know by yourself. Try a lot of N2 worms to get the FEELING!! The same is true for stopping counting. Try N2 and see their typical avoiding behavior. By the way, avoidance behavior of N2 and eat-4 are different. The former reverses as if bouncing off a wall and the latter reverses only after a pause and pause varies from 3-20 seconds. In testing mutants and eat-4 , stop counting at 20 seconds and write down in the assay sheet as 20 seconds.
Tip13. With practice, you can obtain 3 time points/assays from each plate within 25 minutes (in other words, 6 data for N2 and eat-4 and experimental animals). If you find N2 worms take more than 3 seconds for more than 2 out of 6 trials, discard the entire assay.
The above protocol is for the population assay, which tests the multiple worms of the same genetic background. On the other hand, there is the single worm assay, which tests a single worm. The procedure is completely same except that only one worm is transferred to the testing plate. You should take care, however, in the single worm assay that worm often climb up the wall of the testing plate during the assay and is dried up to die. As it is often necessary to recover the worm that has been tested in the single worm assay, take occasionally look at the worm during the assay to make sure the worm is not dried up. If you find the worm on the wall and drying up, use SM buffer to wash it off the wall. However, I would not test the worm once it was washed off by SM buffer during assay.
Paraformaldehyde Worm Fixation Protocol, FIXATION: This protocol is for used to fix worms for staining with antisera. A variation for gluteraldehyde fixation is at the end of this protocol. This is adapted from protocols used in the Li, Horvitz and Kaplan labs.
1)Starve a plate of worms you want to fix for antisera staining (If you want to synchronize for maximal (adult) worm yield/plate).
2) From that plate, chunk to 2 large plates seeded with bacteria. When plates are very full of worms (bacteria is about to be gone, but err on the side of food present starved is usually bad). Wash the worms off with M9 and put them in a 15ml polypropylene tube.
4) Spin the worms in a clinical centrifuge at 2500 RPM for about 30 seconds (or just let gravity-settle for 15 minutes on ice) and transfer the worms in about 1ml to an eppendorf tube.
5) Put the eppendorf tube on ice for a few minutes to cool down the worms (this straightens them).
6) Spin the worms for about 30 seconds (or settle 15 minutes on ice) in an eppendorf centrifuge, remove the M9, and add 500ul ice cold formaldehyde (recipe below).
(Be careful not to inhale paraformaldehyde powder. )
8% pFA in H2O 1.0g paraformaldehyde
(It is easiest/safest to weigh a closed 15ml conical tube, go to the hood, put paraformaldehyde powder in the tube, close it-then weigh the tube again. Adjust water added as appropriate to yield 8% solution. No chance of breathing paraformaldehyde...)
-Heat to 55 to 60 degrees in water bath.
-Heat until the solution clears.
4% pFA in H2O -Add 12.5ml 0.2 M PO4 pH 7.2.
(for use on worms) -Store solution in freezer until needed.
0.2 M PO4 pH 7.2 Buffer: 8.06g KH2PO4
7) Incubate the worms at 4 degrees C for about 24 hours (up to 48 is OK). Longer is preferable with smaller molecules of interest, but reduction time increases proportionally.
8) Wash the fixed worms three times with PBS 7.2.
PBS pH7.2: 16.1g Na2HPO4 . 7H2O
-Adjust the pH to 7.2 with HCl (about 0.9 ml).
-Bring the volume up to 1 liter.
[At this point, the worms can be stored at 4 degrees C to synchronize preparations.]
Beta-Mercaptoethanol Treatment
1) Spin the worms and resuspend them in 1.5ml of
Note that beta-mercaptoethanol is a strong reducing agent, is volatile and toxic .
Or to make 1.5ml [76ul 98% BME
2) Incubate worms at 37 degrees overnight using screwcap tubes to stop the smell of beta-mercaptoethanol from spreading, and set the tubes on the rocker to allow gentle movement of the solution. (For PF/G, rock for 3 days at 37 degrees.)
4) Resuspend worms in 50-100ul of PBS.
[This is another good point to stop if you need to do so.]
1) Transfer about 10ul of worms to a 0.5ml eppendorf tube.
2) Add 400ul of collagenase buffer to each tube.
Collagenase Buffer: 100 mM Tris pH 7.5
1000 u/ml Collagenase Type IV Sigma
3) Incubate at 37 degrees with violent shaking put the tubes in a 50mL Falcon tube and stuff it with paper to the top. Put it in the shaker at 250 rpm.
4) Check the tubes every 30 minutes, starting after 1 hour. When about 20% of the
are broken, stop the reaction by placing the tube on ice.
5) Wash the worms 3 times with PBS.
1) Keep GFP strains in the dark as much as possible to avoid quenching GFP.
2) To do paraformaldehyde/gluteralehyde fixation (PF/G), use 4% pFA and 1% gluteralehyde
(Instead of 12.5ml PO4, reduce by gluteraldehyde volume.)
3) PFA is solid in and stored in the 4 degree fridge. Gluteraldehyde is liquid at room temperature (probably under hood, but may be at 4 degrees in your lab.) Usually ~20% gluteraldehyde. Note that gluteraldehyde is also toxic. Wear gloves.
4) Fixed worms can be stored at 4 degrees in the dark for months.
AMPA assay. This simple protocol was used to assay the effect of AMPA on foraging as mentioned in A.C. Hart, S. Sims and J.M. Kaplan 1995 Synaptic code for sensory modalities revealed by C. elegans GLR-1 glutamate receptor. Nature 378:82-85.
A 7.5mg/ml stock solution of AMPA in water was prepared. 200ul of AMPA stock was mixed with 1.3ml of melted NGM agar (at about 55 degrees C) and poured into a 35x10mm tissue culture dish to yield a 1mg/ml AMPA plates.
An OP50 bacterial slurry was then spread (using a bent pastuer pipet) on the plates and allowed to briefly dry. The OP50 slurry was prepared by spinning down 50ml aliquots of an OP50 liquid culture, then pouring off the liquid and freezing the pellet.
3 to 10 young adult animals were picked to the plates and incubated at room temperature. Foraging and locomotion was scored by two independent observers ignorant of the animals genotype or the presence/absence of the drug at 60 minutes.
1mg/ml AMPA caused hyperactive foraging in N2 animals, but not glr-1(n2461) animals. Neither 0.5 nor 0.25mg/ml AMPA caused reproducible increasese in foraging. Locomotion was not overtly affected.
hart@helix.mgh.harvard.edu
