FREEZING SOLUTION REAGENT

Purpose: Reagent for use in freezing worms

Materials

5. 85 g NaCl

6.8 g KH2PO4 

300 ml glycerol

5.6 ml 1 M NaOH

Milli-Q water to 1 liter

3 ml 0.1 M MgSO4

1. Dissolve NaCl, KH2PO4, glycerol, 1 M NaOH in Milli-Q water with final volume being 1 L. Add to media bottle and autoclave to sterilize.

2. Allow freezing solution reagent to cool before adding the 0.1 MgSO4. Thoroughly mix to incorporate. Store at room temperature.

FREEZING AND THAWING WORMS

Goal: Maintain long-term stocks of worm strains

Materials

Freshly starved worms (no bacterial lawn on plate, worms have not started burrowing)

Freezing solution

1M M9 Buffer

50ml conical tube

Cryovial(s) and cap inserts (5 per strain)

5ml serological pipette(s) (one per strain)

Pipette controller or pipette safety bulb

Foam rack with lid to hold cryovial(s)

Plastic rack (optional to hold cryovials before transferring to foam rack)

Rubber band

Growing Worms to Freeze

1. Place 4-5 L4 worms on fresh NGM with OP50 and grow at 23 °C (grow at lower temperature for temperature sensitive strains). Note: If working with worms that have extrachromosomal arrays select 4-5 worms that contain the extrachromosomal array so that there will be enough worms carrying the array that will survive the freeze and thaw.

2. Allow them to grow until the plate is starved and the majority of the population is arrested at L1. L1s have the best chance of being revived after thawing. The plate will be ready in the second generation.

3. If you are ready to freeze down the worms, proceed to the following steps. If you are not quite ready they can be slowed down and prevented from burrowing by placing them at 16 °C. They can also preemptively be placed at 16 °C if they are close to starving.

Preparing Solution to Freeze Worms

1. Using aseptic technique add 10 ml of the freezing solution and 10 ml 1M M9 Buffer to a 50 ml conical tube labeled Freezing Solution (indicate that it contains freezing solution and 1M M9 Buffer so as not to confuse it with the freezing solution reagent). Invert a few times to mix thoroughly. *You can adjust the volume based on your needs. Just be sure that the two components are roughly 1:1. This solution lasts for about one week, at which point you can discard the solution and retain the conical tube for repeated use to make up this solution.

Washing Worms and Collecting Cryovial

1. Label cryovials and their cap inserts with the strain name. Also, on the cryovial add genotype information. The most important information for the cryovial is the strain name. Make sure it is there. If this is a new strain add the genotype information to the database before proceeding.

2. Place labeled cryovials with their caps off into a plastic rack or a foam rack.

3. With a 5ml serological pipette, aspirate 2.8 ml of the freezing solution (1:1 freezing solution/1M M9 Buffer) and dispense onto plate of worms. Swirl plate. Let sit a few seconds. Swirl again and then tilt plate so that freezing solution and worms pool to one side. Aspirate the worm and freezing solution mixture and dispense 500 μl into each cryovial. Repeat this step for all strains, using a new serological pipette for each strain.

4. Cap all cryovials and transfer them to foam rack if not already there. Place foam lid on foam rack and secure with rubber band and put in -80 °C.

5. Retain the washed plate(s) in case the freeze and thaw is not successful. The few worms left behind will give off progeny and you can chunk or pick from the plate and start the process again.

Thaw Test

1. 24 hours later remove a cryovial from each strain and thaw on bench. This only takes a few minutes. The worms do not like this solution so once thawed immediately move them to a plate. Do not let them sit.

2. Label NGM plates with the genotype. Leave off the strain name as it is not descriptive.

3. Using a sterile glass pipette (Thinking it is best to use sterile glass pipette…sterile…reduce contamination…glass…get maximum amount of worms) aspirate worms from the bottom of the cyrovial (leaving behind as much freezing solution as possible) and dispense along edge of agar, away from lawn. Leave cap vented until the liquid dries and then close lid and turn upside down.

4. The next day check to see if the freeze and thaw was successful. You should see several young worms that have crawled to lawn. However, if they are unc, depending on the severity you may not see this. You should have several young ,live worms nonetheless. In the case of worms with extrachromosomal arrays confirm this under the microscope.

 

GLYCEROL STOCKS FOR BACTERIA

Goal: Maintain long-term stocks of bacteria for worm maintenance and cloning.

Materials (list A if your bacteria is already in liquid culture)

Cryovial(s) (one per strain)

Bacterial strain(s) in liquid culture

50% glycerol (should be cold) (see recipe in Reagents and Solutions)

 

Materials (list B if your bacteria is not yet in liquid culture)

Polystyrene culture tubes with snap caps (14 ml, 17 x 100 mm)

LB agar with appropriate antibiotic (warmed up at 37 °C)

Single colonies of bacterial strains

 

Cryopreservation (list A)

1. Gently swirl culture tube to resuspend bacteria.

2. Add 500 μl bacterial liquid culture to cryovial.

3. Add 500 μl cold 50% glycerol to the same cryovial.

4. Put in -80 °C in an empty cell in our lab stock freezer box of plasmids.

 

Cryopreservation (list B)

1. Streak LB agar with bacterial strain of interest or dispense ~200 μl liquid culture on LB agar (that has appropriate antibiotic incorporated if plasmid has antibiotic resistance gene).

2. Put LB agar plate at 37 °C overnight to grow single colonies of bacteria.

3. Dispense 4ml of LB Broth (with appropriate antibiotic if using bacteria that has plasmid with antibiotic resistance) in culture tube.

4. Load a p10 pipette tip to a pipette and use that to select a single colony. Dispense pipette tip into culture tube. 

5. Put culture tube at 37 °C overnight with shaking.

6. Gently swirl culture tube to resuspend bacteria.

7. Add 500 μl bacterial liquid culture to cryovial.

8. Add 500 μl cold 50% glycerol to the same cryovial.

9. Put in -80 °C in an empty cell in our lab stock freezer box of plasmids. (sketch box with tube entering box)

 

GELS FOR GEL ELECTROPHORESIS

Purpose: Separate DNA fragments based on size and use as a diagnostic test of a digest or extract bands of interest for gel extraction.

Materials

Microwave

Silicone mitt (to handle hot flask)

Kimwipe

1X TAE Buffer (see page x for pdf version and for web version make 1X TAE Buffer drop down and tell user to click to expand and see recipe)

Agarose (we use Goldbio Agarose LE Cat# A-201-1000)

SYBR Safe DNA gel stain

Gel casting stand

Gel tray(s)

Combs

 

Preparing 50 ml of 1% gel (adjust reagent amounts based on your % gel and desired volume)

1. Measure 0.5 g  agarose and add to 250 ml glass Erlenmeyer flask.

2. Measure 50 ml 1X TAE Buffer to flask.

3. Plug neck of flask loosely with Kimwipe to prevent spills.

4. Microwave until completely dissolve. Swirl flask periodically to help agarose dissolve.

5. Add 5 μl to flask and swirl to mix.

6. Add gel tray(s) and comb(s) to casting and pour gel.

 

INOCULATING LB BROTH WITH OP50

Goal: Prepare food source for C. elegans

Materials

NGM plate with contamination-free lawn of OP50

5 ml serological pipette

Motorized pipette controller

LB Broth

Procedure

1. Flame cap and neck of bottle of LB Broth that you’ll inoculate with OP50.

2. Add 1 ml LB Broth to NGM with OP50 and use tip of pipette to scrape OP50 from surface. Aspirate LB Broth onto NGM surface to further dislodge and dissolve OP50. Scrape and aspirate a few times to get an OP50 LB Broth solution.

3. Aspirate OP50 LB Broth solution and dispense in to bottle of LB Broth.

4. Flame neck and cap of bottle. Close bottle, swirl to mix, and leave on bench until it becomes cloudy which is usually after several hours. At the end of the day put at 2 °C for long-term storage. Alternatively, you can put in a shaking incubator to grow the culture quicker.

 

SEEDING PLATES WITH OP50

Goal: Create a food source for worms that localizes them to center of plate so they don’t wander to edge of plate, crawl up the sides, and dry out.

Materials

60 mm x 15 mm plates of NGM (see page x) (if in fridge, leave on bench to bring to room temperature. This will take about an hour)

Pipette controller

10 ml serological pipette(s)

LB Broth inoculated with OP50

Procedure:

1. Stack plates about 8 plates high. (10 ml of bacterial culture seeds about 90 plates)

2. Swirl bottle of LB Broth inoculated with OP50 to resuspend bacteria.

3. Aspirate 10 ml of LB Broth with OP50.

4. Holding the lid of the bottom plate in the stack, lift the other plates and add two drops of LB Broth with OP50 to center of the plate. Note: You can change the diameter of the OP50 lawn that the worms will crawl on by changing the distance between the pipette tip and NGM surface. The shorter the distance, the smaller the lawn. The larger the distance, the larger the lawn.

5. Repeat step 4 for every plate you seed until you have seeded all the plates in the stack.

6. If you dispensed everything in your pipette and still have plates left to seed. Get a new serological pipette to aspirate the bacterial culture. A pipette should never enter the bottle twice as this can introduce contamination.

7. Leave plates on bench for a day to dry and then move to 2 °C for long-term storage.

MAKING SLIDES FOR COMPOUND SCOPE

Goal: Temporarily paralyze worms and image using compound scope

Materials

2% agarose (melted)

1 mM levamisole

4 microscope slides (25 x 75 x 1.0 mm)

Labeling tape or masking tape

200 μl pipette

Cover glass (22 x 22 mm)

Making agarose pad:

1. Place two strips of tape directly on top of each other on two of the microscope slides. Orient the two slides with tape and a third slide horizontally on your microscope with their long edges touching top to bottom. The slide without tape should be the middle slide. (The slides should be on the same plane and not stacked on top of each other). The thickness of the tape layers determines the thickness of the agarose pad. Orient the fourth microscope slide vertically and make its long edge flush with the short, right hand edges of the other 3 slides.

2. Dispense 70 μl agarose in the center of the middle slide and place the vertical slide on top of the agarose to flatten it (such that it lays across all three slides). After a few seconds, carefully slide this microscope slide away to uncover your agarose pad. Note: Bubbles may form when you make your agarose pad. If you feel there are enough present to impair imaging you can wipe the agarose pad off the slide with a kimwipe and start over. You have now created your agarose pad. Retain the microscope slides with tape to make more slides.

3. If screening worms, add 2 μl 1 mM levamisole to the center of the pad. This amount reduces the spread of worms and thus decreases imaging time. For all other imaging use 3-4 μl of 1 mM levamisole.

Transferring worms to agarose pad

1. Working quickly and using aseptic technique put bacteria on worm pick and select worms of interest. They will stick to the bacteria. Touch your pick on the surface of the levamisole. Try not to go down to the pad. Gently wiggle pick in levamisole to dislodge worms. Continue this step 1 until you have gotten all your worms of interest. If the levamisole drop starts to dry, add more in 2 or 3-4 μl aliquots. As the levamisole dries the worms will clump together which may make imaging difficult depending on what you are imaging.

2. Slowly place coverslip over pad starting with one edge and carefully placing it down while trying to avoid the creation of air bubbles. (Imagine closing a book cover as opposed to placing the whole book down). You can wait until worms stop moving or image them right away.

 

STOCK SOLUTION OF 10 mM

Purpose: use as a paralytic agent to significantly reduce movement of worms during imaging

Materials

Levamisole (Acrōs Organics)

1 M M9 Buffer

2 ml microcentrifuge tubes

50 ml conical tube

Procedure:

1. Measure 0.21 g levamisole into 50 ml conical tube.

2. Add 1 M M9 Buffer to 50 ml.

3. Dissolve by vortexing.

4. Aliquot into 2 ml microcentrifuge tubes and store at -20 °C.

 

WORKING SOLUTION OF 1mM LEVAMISOLE

Purpose: For imaging with compound scope

Materials

10 mM stock solution levamisole

Water or 1 M M9 Buffer (to dissolve levamisole)

1.5 ml microcentrifuge tube

Procedure:

1. Add 90 μl water or 1 M M9 Buffer to microcentrifuge tube. Note: If you dilute in M9 Buffer you have to put it in the fridge or else it will grow bacteria relatively quickly. If it is diluted in water bacterial growth takes longer so you can leave it on the bench.

2. Add 10 μl of 10 mM levamisole to the microcentrifuge tube. If the 10 mM levamisole is frozen you can quickly thaw by rubbing between palm of hands.

3. Pipette up and down or vortex to mix.

 

2 L LB AGAR (adapted from WormBook)

Materials

20 g tryptone

10 g Bacto-yeast

10 g NaCl

30 g agar

6 L glass Erlenmeyer flask (size can vary but should be at least twice the volume of media)

100 mm petri dishes

Procedure:

1. Add stir bar, 20 g tryptone, 10 g Bacto-yeast, 10 g NaCl, 30 g agar and Milli-Q water to 1 L to Erlenmeyer flask. Autoclave to sterilize.

2. Using aseptic technique dispense 25 ml of agar per plate.

3. Leave at room temperature to dry and the following day store at 2 °C.

 

2 L LB AGAR WITH AMPICILLIN (adapted from WormBook and Cold Spring Harbor protocols)

Materials

20 g tryptone

10 g Bacto-yeast

10 g NaCl

30 g agar

6 L glass Erlenmeyer flask (size can vary but should be at least twice the volume of media)

2 ml of 100 mg/ml ampicillin (aliquots in fridge or thaw from -20 °C)

100 mm petri dishes

Procedure:

1. Add stir bar, 20 g tryptone, 10 g Bacto-yeast, 10 g NaCl, 30 g agar and Milli-Q water to 2 L to Erlenmeyer flask. Autoclave to sterilize.

2. Put LB agar on stirring plate with stirring on and when LB agar has cooled to 50 °C add 2ml of 100 mg/ml ampicillin and continue stirring to mix thoroughly.

3. Using aseptic technique dispense 25 ml of LB agar-ampicillin into each 100 mm petri dish.

4. Leave at room temperature to dry and the following day store at 2 °C.

 

COMPETENT CELLS

Goal: Create cells that can be transformed with plasmids of interest. This protocol starts with competent cells. To get a stock of competent cells you want them to divide, however cell division causes them to lose their competency so this protocol makes these newly divided cells competent.

Time: 3 days. On the third day if you were successful you can confidently use these cells in your experiments.

IMPORTANT: Try to keep things as sterile as possible. The competent cells don’t have any antibiotic resistance. You don’t want any bacteria growing with them and getting transformed later on. Also, be gentle with the cells throughout this protocol. Any roughness kills the cells.

Materials

50% glycerol (cold)

100 ml 100 mM CaCl2 (cold)

100 ml Milli-Q water

10 ml of 85 mM CaCl2, 15% glycerol (freezing solution)

Competent cell freezing solution (see below)

Competent cells (Can start with cells from a kit or with ones that you have successfully made. We initially started with TG1 cells from Zymo’s Mix & Go kit)

200 ml LB Broth (100 ml in two separate flasks)

Two 14 ml culture tubes with snap caps (17 x 100 mm)

Cuvettes

Spectrophotometer

Two 50 ml Oakridge conicals (cold)

10 ml serological pipettes (cold)

25 ml serological pipettes (cold)

Sterile 1.5 ml microcentrifuge tubes (cold, to store aliquots of competent cells)

Ice

Two ice buckets

Thaw competent cells:

1. Thaw TG1 cells on ice. This should take about 10-15 min. You will notice that a pellet will settle out/form. Gently tap the bottom of the tube to re-distribute the cells. Remember to be gentle!

Competent cells overnight culture (start this the night before):

In the hood:

1. Using a 10 ml serological pipette and aseptic technique. Put 5 ml of LB broth into 2 separate culture tubes.

2. With a flame going nearby, use a sterile filtered 20 μl tip to add 10 μl of the competent cells to one of the LB broth culture tubes and sterile filtered 200 μl tip to add 90 μl to the other LB broth culture tube. Put the caps on loosely for air circulation. We are starting with two different concentrations so that in subsequent steps if we miss the mark we have a back up.

3. Put in a 37 °C shaking incubator for about 16-18 hours.

2nd culture of competent cells:

1. Inoculate two separate 100 ml of LB broth with 50 μl of the overnight cultures. One inoculation per flask. Put these flasks and another 100 ml LB broth to serve as a blank in a 37 °C shaking incubator.

2. You want to stop the cells while they are in log phase. This is a good stage to try to make cells competent.The O.D.600 that you should aim for is 0.4. It will take about 1.5 hours to reach this O.D.600 but you’ll need to check in increasingly shorter intervals as you get close to that O.D.600

Measuring O.D.600:

1. Remove all three flasks from the shaker and bring to the hood and add some of the LB broth to a separate cuvette. Be careful when handling the cuvette. You don’t want any skin oils to touch the area where the light passes through.

2. We use a Beckman Coulter DU 730 Life Science UV/Vis in Bryen Jordan’s lab.

3. Choose the “Fixed wavelength setting” at 600 nm.

4. Insert your blank into the instrument and select “Blank” and then “Read”. The “abs” or absorbance is the O.D.600

5. Measure your two cultures and record the values. These values will help you project when you might be getting close to an O.D.600 of 0.4. The upper limit is 0.6. If you overshoot that, you will need to start another overnight culture.

Preparing Refrigerated Centrifuge:

1. We use the Beckman Coulter Avanti J-E centrifuge in the Molecular Biology Core in room 328. The refrigerated rotor JA-20 that is stored in the 4 °C fridge in the same room.

2. Use the cart in the room to transfer the rotor to the centrifuge. Insert the rotor and screw down the lid.

3. Set centrifuge to spin at 3000 rpm at 4 °C to get the chamber cool for when we are ready to spin our cells down. The rpm is not so essential for prepping the chamber, but the temperature is.

Spin down cells:

4. Put the 50 ml oakridge conicals on ice and put the lid on for insulation. Whichever, overnight culture reaches an O.D.600 of 0.4 first spilt the culture in half between the two oakridges.

5. Let them sit on ice for about 20 min.

6. Centrifuge at 5,000 rpm for 15 min. at 4 °C.

Wash cells:

1. Back in the hood pour off the supernatant and using a 25 ml serological pipette resuspend cells in 20 ml of cold 100 mM CaCl2. Use one serological pipette per tube. To resuspend you can 1)swirl or 2)pipette up and down or 3)gently vortex. We have successfully vortexed (7.25 speed) and also pipetted up and down. Remember again to be gentle, many things can kill the cells.

2. After resuspending, put the back on ice, close lid and let incubate for 1 hr.

Spin Down Cells:

1. Before spinning down cells, put your 1.5 ml microcentrifuge tubes on and your freezing solution on ice.

2. Centrifuge cells at 5,000 rpm for 20 min.

3. Using a serological pipette remove the supernatant. The pellet will be a little scattered. Scattering is normal and you will lose some of the pellet in this step. Try to get all the supernatant out. If a very tiny amount is left behind that is fine.

Freezing cells:

1. Add 2 ml of the freezing solution to each tube and use the serological pipette to pipette the solution up and down to dissolve the pellet.

2. Add 100 μl of the cells in freezing solution to each 1.5 microcentrifuge tube.

3. Snap freeze them with liquid nitrogen.

4. Store in our competent cell freezer box at -80 °C.

Testing competent cells:

1. Transform cells with a plasmid of your choice to make sure you were successful in making competent cells. In the event that the cells are not competent, it would be a let down and time sink to wait until your experiment or someone else’s to find out.

RESUSPENDING PRIMER(S)

Materials

Primer(s)

Micro/minicentrifuge

TE Buffer

50 ml conical tube

1000 μl pipette

Spin Down Primer(s)

Primers are usually shipped and arrive in a lyophilized state. During shipping the flakes or pellet can move around. If you don’t centrifuge it first the primer may pop out when you open it!

1. Centrifuge for about 1 minute. You should see a thin film ring at base of tube after centrifuging.

Resuspend in TE Buffer

1. Add TE Buffer to a 50 ml conical tube to keep as your working stock.

2. Add appropriate amount of TE buffer to primer(s) as indicated on documentation that accompanied it. The final concentration will be 100 μM. Be careful: Once you add TE Buffer to a primer, get rid of the pipette tip immediately so you don’t forget and then cross-contaminate your primers.

3. Vortex to mix. Add name of primer to cap to help with quick identification. Store at -20 °C in a cardboard freezer box.

NANODROP

Goal: Quickly and easily quantify and assess purity of DNA sample(s)

Materials

DNA sample(s)

Milli-Q water

50 ml conical tube

1. 5. μl pipette

Kimwipes

Gloves

Clean NanoDrop Prior to Use:

1. Add Milli-Q water to conical tube and bring that and all the other materials to the NanoDrop instrument.

2. Wearing gloves, wet (but not dripping wet) Kimwipe with Milli-Q water and wipe down stage, surrounding area and area that touches stage on lid and surrounding area. It is important to wear gloves so that you avoid getting natural oil from skin onto NanoDrop stage.

Blank NanoDrop:

1. Add 1.2 μl Milli-Q water to middle of stage and select “Blank” in the software.

2. Wipe stage with Kimwipe and add 1.2 μl Mill-Q water to stage. Enter sample name “Milli-Q Blank” or something similar and select “Read” in software. Wipe with Kimwipe and proceed to measure first sample.

3. Before measuring a new sample, enter the sample name, wipe the stage and surrounding area, and part of lid that touches stage and surrounding area with a Kimwipe that is wet with Milli-Q water. Add 1.2 μl of sample. Dispose of pipette tip immediately. Select “Read” in software. Record the value either by writing down or taking a picture.

4. At some point the software may prompt you to save your data file. Save in an appropriate folder. We have a main for Kurshan Lab as this is shared equipment. Record the concentration and the 260/280 value. The 260/280 value is an indicator of the purity of nucleic acids. It is the ratio of absorbance at 260 and 280 nm. The amount of light absorbed at 260 nm is proportional to the nucleic acid concentration of the sample. Proteins (which are considered contamination in these samples) absorb at 280 nm. A ratio of ~1.8 is considered “pure”. 

5. Do a final wipe down of the NanoDrop as described above.

WORM CROSSES

Purpose: To generate a genotype of interest

Important: Draw out your crosses before starting and do not skip any steps. It is not worth the time that will be wasted due to poor planning. Figure out what you need and when you need it i.e. the different strains and sexes of worms. You may need to synchronize worms at the start of our crosses that you will use at a later stage in your crosses. Doing this saves you time.

Materials

~4 L4 hemaphrodite C. elegans

20 (at least 10)  male C. elegans (per cross) (use more if the cross has a low success rate)

1 -2 60 mm NGM plates (per cross) without any bacterial lawn (the extra plate is in case something goes wrong)

LB broth with OP50

1 ml serological pipette

Pipette controller

1 mM levamisole (Optional, depending on difficulty of cross)

Setting up cross plates:

1. About an hour before you plan to set up your cross(es), seed your NGM plate(s) with OP50 LB broth. Put one drop close to the NGM surface. You want the lawn to small and thin to increase the chances of mating.

Basic setup of cross:

1. Place your L4 hemaphrodites on your cross plate. Move males that you will use for mating to a “checkpoint” plate and let them crawl around. This makes it easier to avoid carrying eggs or even hermaphrodites to the cross plate if the source plate is super crowded. Eggs and unwanted hermaphrodites can quickly destroy your crosses. Place cross plates at 23 °C, if they are not temperature sensitive. Otherwise, put them at 20 °C. Some important considerations when setting up cross:

1. If the males that will be added to the plate are pretty unc or having other mating defects, paralyze the hermaphrodites with levamisole on a separate plate and then move them to the cross plate.

2. Select the youngest males you can identify. Virility decreases with age. Never use starved or 3-4 day old adult males for crosses.

3. Make sure that all the strains you will be using in subsequent steps are grown at the same temperature as your initial cross so that they are synchronized.

Cross Maintenance:

1. Keep track of your crosses. Write out the day you completed each step or pre-write the date you plan to do each step and check off each step as you complete it.

2. The final genotype of desired cross progeny may be the result of several crosses. Each day setup a new male cross of desired genotype or chunk from a plate of almost starved males so you always have males ready for the next step(s) in your cross. Any hermaphrodites that will be used in later crosses should also be maintained so they are ready when you need them.

3. Check early on to see if your cross worked. If you see male progeny your cross was successful.

4. Select cross progeny based on your plan and set up additional crosses as needed.

Obtaining desired cross progeny:

1. Once you have validated your cross, freeze down worms as soon as possible. Mistakes happen and if somehow a mix up happens or your strain gets completely lost you will have to redo the cross. For example, you may contaminate your plate with non-target cross progeny. If you weren’t paying enough attention to notice the different genotypes on the plate, over time and with regular chunking or picking you may inadvertently lose the strain of interest. 

BLEACHING WORMS TO REMOVE CONTAMINATION OR ”SYNCHRONIZE” WORMS

Goal: Get rid of contamination early before it becomes a persistent and widespread problem or to (almost) synchronize worms for crosses or other purposes.

Materials

Gravid adults

Fresh 60 mm NGM plate with OP50 lawn

15 ml conical tube wrapped in aluminum foil

Bleach (once opened it starts to go “bad” over time, so make sure you are pulling for a batch that is still working)

1 M NaOH

50 ml conical tube

1. 5. ml microcentrifuge tube

Preparing bleaching solution:

4. Add bleach to the 15 ml foil-wrapped conical tube. Properly label the cap and the foil.

5. Add 1 M sodium hydroxide to a properly labeled 50 ml conical tube.

6. Add bleach and 1 M sodium hydroxide 1:1 in a 1.5 ml microcentrifuge tube.

7. Note: Keep the 15 ml and 50 ml stocks of bleach and 1 M NaOH, but make the bleach solution fresh the day you use it.

Bleaching Worms

1. Label the NGM plate with the genotype of the worms to be added and also “(b)” to indicate that it is a bleach plate. Add 10 μl of the bleaching solution to the edge of the plate. It should not be near the lawn. You want the L1s that hatch, to crawl away from the spot of bleach solution and onto the lawn.

2. Add about 20 gravid (egg containing) adults to the drop of bleaching solution. Pick adults that have at least two rows of eggs. You don’t want adults that just started producing eggs. The worms that just started producing eggs will 1) have less eggs and 2) will likely contain eggs early in development that will not survive the bleaching. The agar beneath the solution is softer than normal, so be careful not to cut through the agar when you are adding worms.

3. You want the bodies to break down, only leaving eggs behind. If you notice the drop is drying up before this happens add the bleaching solution in 10 μl aliquots at a time.

4. You can leave the petri dish cap off or partially off to allow the solution to dry. Once dry, close the lid.

5. The next day move L1s to fresh NGM with OP50. They are not fond of the bleach solution.

50 ml 5 mg/ml CHOLESTEROL STOCK SOLUTION

Purpose: Component of NGM

Materials

125 mg cholesterol

100% ethanol to 50 ml

0.2 μm nylon syringe filter

Syringe

Sterile 2 ml microcentrifuge tubes

Procedure:

1. Add 125 mg cholesterol to 50 ml conical tube. Add 100% ethanol to 50 ml. Put in 42 °C water bath and the vortex. Repeat these two steps until cholesterol is fully dissolved.

2. Filter sterilize the cholesterol solution in the 2 ml aliquots per microcentrifuge tube. Store at 2 °C for short-term use and -20 °C for long-term use.

MICROINJECTION PADS

Purpose: Surface to place and rescue worms from for microinjections

Materials

2% agarose (Goldbio Agarose LE)

200 μl pipette tips

24 x 60 mm # 1 Richard Allen Scientific Slip-Rite cover glass

Fine tip permanent marker

Razor blade or scissors

Prepare pipette tips to dispense agarose:

1. Cut the bottoms of several 200 μl to use for aspirating and dispensing agarose. Your style of tip will determine how much you need to cut off. You want to cut off enough so that you can aspirate enough agarose and also be able to dispense it.

Making agarose pads:

1. Aspirate 100 μl 2% agarose and dispense onto center of cover glass. Quickly cover the agarose with another cover glass and press down on both ends of cover glass and center (if needed) to flatten and spread out the pad. Immediately slide cover glasses away from each other. Whichever one has the agarose pad attached to it, write “UP” in the right hand corner of the same side of the cover glass as the pad.

2. Bake them in an incubator overnight at 65 °C.

3. Store them in the original cover glass container.

2L NEMATODE GROWTH MEDIUM (NGM)

Purpose: Medium that C. elegans are maintained on in the lab.

Materials

6 g NaCl

34 g agar

5 g peptone

1,950 ml Milli-Q water

2 ml 1 M CaCl2

2 ml 5 mg/ml cholesterol

2 ml 1 M MgSO4

50 ml KPO4

60 mm petri dishes

Three 2 ml serological pipettes

One 50 ml serological pipette

Milli-Q water (to flush tubing after dispensing media)

Labeling tape

Autoclave tape

Peristaltic pump (we use Integra MEDIAJET)

Tubing and filling nozzle (specific to our system)

Preparing NGM:

1. Mix 6 g NaCl, 34 g agar, 5 g peptone, and 1,950 ml Milli-Q water with a stir bar in a 6 L Erlenmeyer flask. Apply a double layer aluminum foil cap to mouth of flask and then slightly vent it. Fill a glass media bottle with about 300 ml of Milli-Q water to flush tubing once done dispensing media. Label both containers. This is especially important when using a shared autoclave facility. Apply autoclave tape to both containers. Autoclave to sterilize. With our particular system we put the flask and bottle in an autoclave safe tub with some water. Run the autoclave for 53 min. on the liquid cycle at 121 °C. At the same time sterilize your tubing and dispenser.

Sterilizing tubing to dispense media:

1. Sterilize the tubing and filling nozzle at the same time you are sterilizing the NGM. Wipe down tubing and dispenser with Kimwipe to remove any dust, debris, dried agar and then wrap in a coil. Wrap both ends of tubing with foil so that about 6 inches on each end is cover. Cover entire coil of tubing in two layers of foil and put a small piece of autoclave tape on the outside. Autoclave on the liquid cycle for 35 min. with a 20 min. dry time.

Loading carousel with petri dishes:

1. Turn on MEDIAJET and select the LOAD CAROUSEL option. Remove dishes already in carousel if needed.

2. Time saving tip: Instead of loading plates a handful at a time, place entire sleeve in column and then cut off bottom of sleeve and also up along the side. Pull sleeve up and away.

3. Take care that the stacks of dishes in any column do not go beyond the notch on the rods of the columns. This will cause the instrument to jam and malfunction during operation.

4. At least one column in the carousel should be free before turning on the MEDIAJET. This ensures that there is enough space to load dishes to be filled and then stack them once filled.

Adding post-autoclave reagents:

1. Place autoclave tub with flask on a stirring plate and turn on stirring. Using an infrared thermometer check the temperature and start adding the post-autoclave components when the temperature reads 74 °C (if using a water bath wait till is 55 °C). Check the temperature by placing the thermometer directly above the opening of the flask so laser points straight down and is one place. There shouldn’t be two “dots”.

2. Using aseptic technique and the serological pipettes (one per reagent) add 2 ml 1 M CaCl2, 2 ml 5 mg/ml cholesterol, 2 ml 1 M MgSO4, and 50 ml KPO4.

3. If adding any antibiotics add them now.

Selecting program in MEDIAJET:

1. Go to the MAIN MENU and then press FILL DISHES and select 01 PRG 01 or the first program in the list in case at the time of reading this document the name has changed. Check the PROGRAM SETTINGS to make sure that it is for 60 mm dishes and that the Dispensing volume [ml]: is 12 ml.

2. Now press START PROGRAM.

The instrument should load the first place into the turnstyle that is under the cover and in line with the filling nozzle.

Connecting setting up tubing and filling nozzle:

1. Carefully unwrap foil and using aseptic technique retract filling nozzle inside if it is extended and insert into the MEDIAJET filling nozzle mounting. Now fully extend the filling nozzle.

2. Carefully place the other end of the tubing inside of the flask with the NGM and secure foil cap over opening of flask.

3. Open pump cover and insert tubing such that there is about 30 cm (12 in.) stretch of tubing between filling nozzle mounting and pump. Tubing should be flush with back wall of pump. Close cover.

4. Important: Check to make sure the filling nozzle is fully extend. If you forget this part agar will spill everywhere!

Priming tubing and filling dishes:

1. Hold down “Prime” button to completely fill the tubing. You’ll know tubing is completely filled when you see media being dispensed into the petri dish.

2. Press START.

Clean-up:

1. Remove filling nozzle from MEDIAJET and flush with the sterile Milli-Q water, catching flush water in some sort of “waste” container. Pour flush water in sink. Wipe off any agar that might be on the filling nozzle.

2. Clean up any agar that may have spilled in the housing of the MEDIAJET.

Plate storage:

1. Leave plates in carousel until the agar has solidified. Once solidified move to bench leave on bench for about 3 days to monitor for signs of contamination. You can also speed up this process by placing a few of the plates at 37 °C. Most contamination will present itself after 24 hours at 37 °C. Store plates upside down at 2 °C.

TRANSFORMATION

Purpose: Have competent cells take up plasmid of interest. These transformed cells can be grown up and the plasmids can be extracted for cloning and microinjection.

Materials

Competent cells

Ice

Ice bucket

LB broth

50 ml conical tube

LB agar with ampicillin

Rattler plating beads

42 °C water bath

Shaking incubator

Warm up LB agar ampicillin plate(s):

1. The last step will be to plate the transformed cells onto LB agar with ampicillin and grow up single colonies at 37 °C. Label the plate(s) and put at 37 °C to warm up.

Gently thaw competent cells and plasmid(s) on ice:

1. Remove competent cells from -80 °C and put on ice and label tube(s) with name of plasmid that it will be transformed with. If transforming a cloning product you should also pull out an additional competent cell tube to serve as a control. Put the plasmid(s) on ice as well (those don’t need to thaw gently or on ice. If not thawed,they just need to be thawed in general.)

2. Starting with this step and after each subsequent step gently flick the competent cells tubes.

Incubate competent cells with plasmids on ice:

1. Add 50 μl of plasmid to the thawed competent cells and incubate on ice for 30 min. The timing here is important. If you are looking to save time you can do so during the recovery step, not this step.

Heat shock, recovery, and plate:

1. Using aseptic technique add ~50 ml LB broth to a sterile 50 ml conical tube and keep on bench to use whenever you are doing recoveries.

2. Heat shock the competent cells in a 42 °C water bath for 1 min. and then put back on ice and add 200 μl LB broth.

3. Tape tube of cells to base of shaking incubator and incubate at 37 °C for 45 min.

4. Add 15-25 rattler plating beads to LB agar ampicillin plate and add respective cells to plate. Close lid and swirl plate keeping it horizontal to the ground. Dispose of rattler beads into bottle of ethanol with the other waste rattler beads. The beads can later be sterilized and re-used.

5. Put plates upside down at 37 °C overnight. If your transformation was successful, wrap the LB agar ampicillin plate with parafilm and store at 2 °C.

GEL EXTRACTION

Purpose: To extract fragments of DNA from a digestion gel that will later be used in cloning.

Follow the instructions provided with the gel extraction kit you are using. We are currently using QIAquick PCR and Gel Cleanup Kit. Add the following steps to your gel extraction process.

1. Keep a working stock of isopropanol on your bench. It is light sensitive so wrap a 50 ml conical tube with foil before dispensing into tube. Be sure to probably label the tube itself, the foil, and the cap with the date that original isopropanol bottle was opened and when you created this working stock.

2. 0.4 g is the max amount of gel a column can take. Round samples over 0.100 g up to 0.200 g.

3. Since the gel purification columns can only take 750 μl at a time, you may have to repeat Step 3 a few times depending on your sample size. It is okay to run a column repeatedly to balance another column that had more sample.

4. Hold the waiting step for elution for 1 min. and then centrifuge for 1 min.

5. At the very end when eluting, cap the columns with the 1.5 ml microcentrifuge tubes that you will use to collect your DNA. In case the cap breaks it is more important to have the microcentrifuge as opposed to column cap on.

EMS MUTAGENESIS

Day -4: Prepare 6-10 plates with 4 or 5 L4 animals per plate and put in 20°C incubator. 

Day 0: Mutagenesis:

1.      Prepare a solution of 0.1 M NaOH, about 500 ml in a plastic beaker. This will be used to neutralize the EMS from pipette tubes, tubes. LABEL THIS CONTAINER with your name, the date and “SODIUM HYDROXIDE”.

2.      Wash each plate with 1 ml M9. Spin down gently in picofuge.

3.      Collect worms in 15 ml conical tube using glass pipettes. Have a final volume of 2 ml in this tube.

4.      Add 2 ml of M9 into another 15 ml conical tube. Add 20 μl of EMS into this solution.

5.      Vortex EMS-M9 tube until EMS dissolves.

6.      Pour the 2 ml solution of EMS-M9 into conical tube with worms.

7.      Put on rotator for 4 hrs wrapped in foil (EMS is light sensitive).

8.      Add 10 ml of M9 to tube. Spin down gently using centrifuge in hood. Aspirate supernatant with glass pippetes. Repeat 5 times.

9.      Suck up worms using a glass pipet and plate onto seeded plates. Let the worms rest for 2 hrs.

10.   Pick mid to late L4s onto new plates (5 per plate, ~10 plates). These are your P0s.

11.   Age the animals 24 hrs at 20°C after the end of EMS treatment. This will allow you to avoid the first eggs which may have already been meiosis at the time of mutagenesis.

Day 1: Transfer P0s

12.   Pick P0s onto a new plate and put back at 20°C.

13.   Dispose of pipette tips, tubes, in normal trash 24 hrs after EMS mutagenesis. Dump 0.1 M NaOH down the sink.

Day 4:

14.   Pick F1s: Put 2-3 animals per plate, ~30-90 plates (start with fewer, ramp up if you can).  You can either pick all L4s and divide into three batches: 16/20/23 °C, or divide by picking 1/3 L3s, 1/3 L4s, and 1/3 YAs and put them all at 20 °C.

Days 7-9+:

15.   Score F2 L4s. Check ~16 animals per number of F1s that were on the plate. Rescue any potential mutants and single them onto new plates placed at 20 °C.

Days 11+:

16.   Score F3s to check for transmission of phenotype. Keep everything at 20 or 16 °C to prevent loss of sickly strains.

INJECTIONS

Prepare 10ul of injection mix:

• • -DNA for injection at appropriate concentration (usually 1-10 ng/μl for most transgenes)

  • -Coinjection marker (50-100 ng/ μl)

  • -Milli-Q water to 10 μl

1. Spin down for 10-20 min. Carefully take top 8 μl into a new tube. Spin again for another 10 min.

Preparing needle for injection:

1. Pull needles (program #?). let the puller warm up for a few minutes before pulling the first needle, or pull a few “blanks” (no glass) first. Pull a 2-3 needles per injection mix so you have extras. store vertically in the glass jar.

2. Lay one injection needle on top of a plate cap on the dissecting scope with both the tip and back suspended off the plate cap.

3. Carefully take 0.5 μl from the top of the injection mix and pipette onto the back of your injection needle. Let the needle fill by capillary action, then try to remove any air bubbles by flicking and/or shaking the needle.

4. Remove the needle holder from the stage clip, then untwist to remove the head of the needle holder and take the previous needle out (save it for making a worm hook later). Push the new needle into the needle holder head so that the back of the needle sticks out by about ½ cm from the back of the needle holder head. Screw the head with the needle back onto the needle holder and place it back onto the stage clip (tighten the screws if necessary).

5. Turn on the injector (with the tubing removed), let it come up to pressure before inserting the tubing (it twists and clicks in).

6. If you need to make a new needle-breaking coverslip:

Put a large drop of mineral oil on the center of a fresh blank cover slip (no agar pad).

Take a new capillary tube and hold it over the burner flame pulling on both sides until the tube elongates forming a thin filament. Break the filament off one side of the capillary tube and lay it down on the coverslip so that it sticks to the mineral oil, positioned perpendicular to the long axis of the cover slip. Break the filament off the send capillary tube so that only the filament is left on the coverslip (you can trim any extra pieces off with scissors).

7. Place the needle-breaking coverslip on the stage and bring it into focus, positioning the glass strip vertically (first using the low mag then the high mag objective). 

8. Position the needle tip as close as possible to the cover slip by eye, then, using the low mag objective, bring the needle tip into the center of the field of view using first the coarse manipulator (sweep it vertically back and forth to try to see the shadow) then the fine manipulator. Once it is in view/~in focus in the center of the field of view, switch to high mag and re-center/bring it back into focus (using the fine manipulator, not the focus knob). Bring the needle close to the glass strip and then slowly ram it into the glass strip, checking each time by clicking the right mouse button to see how large/fast the bubbles come out (bubbles should come out rapidly). Once you are satisfied with the break size of the tip, lift the needle slightly by turning the vertical fine manipulator by a few twists. 

Preparing the worms:

1. Select gravid young adults with 5-10 eggs and pick them onto a new plate (fewer eggs = more injected progeny, but harder to inject).

2. Take an injection pad from the box and place it agar side up on top of a plate cover. Add a small drop of mineral oil to the side of the agar pad. 

3. Make sure your worm pick is smooth and flat to prevent worms crawling up.

4. Use your pick to put a small, ~2x worm-sized drop of oil onto the agar pad.

5. Move a worm from the bacterial lawn to the side of the plate (outside the bacterial lawn). Let it crawl away from the residual bacteria.

6. Dip the bottom surface of your pick into the large spot of oil to get a drop of oil onto the bottom of your pick, but try not to get any oil onto the top surface of your pick. Use the oil to pick up your worm (you might need to “swipe” at it). 

7. Place your worm onto the oil spot on the agar pad, try to get it off your pick quickly but not too roughly. The worm will stick to the pad- try to get it to stick straight not curled. (If you are putting more than one worm down, try to align them with their dorsal sides facing roughly the same way.)

8. Check your needle again to make sure it hasn’t clogged up since you last checked it (re-break if necessary). 

9. Raise the needle again, and slide the breaking coverslip out of the way. 

10. Switch to low mag and slide the worm pad coverslip into position, positioning the worm vertically with dorsal side facing the needle. Center the worm and switch to high mag. Focus on the worm and then bring your needle back down into focus (using the fine manipulator). 

11. Position your worm (by moving either the coverslip or the round stage) so that you can see one of the gonad arms. Focus up and down until you see the middle of the gonad, which has a “speckly” appearance (like dark grains of sand). Bring your needle tip into the same focal plane using the fine manipulator. Push the needle in until it punctures the cuticle and position the needle tip in the center of the gonad arm. Inject by pushing the right mouse button and look for the solution “wave” to course up and down the gonad arm, ideally rounding the corner towards the ventral side of the gonad arm. If instead of the vertical wave you see what looks like a bunch of grapes, your needle was positioned outside the gonad. Retract, refocus/reposition, and try again. After injecting one gonad arm, move the worm and inject the second gonad arm (anterior/posterior).

Recovering injected worms:

1. When done injecting, raise your needle slightly with the fine manipulator, slide the coverslip out, place it back on the dissecting scope, and add a drop (~1ul) of M9 onto your worm. It should slowly become unstuck from the pad and start thrashing a bit.

2. Make a hook from the old injection needle you had saved by passing it over the top of the ethanol burner flame until the tip rounds up into a hook. Use the hook to pick up your worm(s) from the M9 and place them onto a new plate (right next to the food), being careful not to puncture your plate with your hook. Add ~3 worms per plate and let them recover at 20deg.

Checking progeny for coinjection marker: 

1. Check them after 3 days for F1s with the coinjection marker. Single F1s (~24 L4s for a regular transgene injection). Check them after 3 days (at 23deg) for F2s that have the coinjection marker (~25-30% of your F1 plates will have some F2 progeny with the marker). Save the plates that have some F2s with the marker and assess them for phenotype.

Changing needle (when needed):

1. If you need to change your needle, push the “menu” button and then the “change capillary” button on the injector. Change your needle and then push the menu button again to get back to the injection screen.

Proper shutdown of injector at end of session:

1. When you are done injecting, either push “standby” once if someone is going to inject after you, otherwise push and hold the standby button to release the pressure, then turn off the injector and remove the tubing. Injector instruction booklet is there if you need to consult it.