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Protocols for worm media and maintenance

Most of the protocols can be found on wormbook. The ones below are from BioProtocols. He, F. (2011). Common Worm Media and Buffers. Bio-101: e55. DOI: 10.21769/BioProtoc.55.

Before using any of these protocols, please cross-check with other sources. recalculate the given weights with the molar weights on our chemicals. When you verified a protocol, make a note 'verified by [NAME]' under the headline.

Stock solutions

  • Cholesterol 5 mg ml-1 in 95% EtOH
  • 1 ml 1 M CaCl2
  • 1 ml 1 M MgSO4
  • 25 ml 1 M potassium phosphate (pH 6)
  • Nematode growth medium (NGM) liquid

1M potassium phosphate (pH 6)

Alternatively: Make the following two solutions separately, then mix to obtain correct pH (6.0):

A. For 250 ml 1M KH2PO4 (monobasic): Add dH2O to 34.0 g KH2PO4 until final volume (250 ml) is obtained

B. For 200 ml 1M K2HPO4: Add dH2O to 45.6 g K2HPO4 *Make sure the salt in the solutions is completely dissolved.

  1. Add K2HPO4 solution to KH2PO4 solution to bring the pH up from 4.0 to 6.0 (will take about 100 ml of K2HPO4).
  2. Autoclave 15 min. liquid cycle
  3. Store at room temperature.

Alternatively: For 1 liter, dissolve 136.1 g KH2PO4 in about 800 ml dH2O, then adjust to pH 6.0 with solid KOH (approx 15 g) before bringing up to volume. Make 100 ml aliquots and autoclave. Jun's note: you may also use ~53ml of 5M KOH solution.

Nematode growth medium (NGM) agar

For the maintenance of worms on plates. For liquid NGM just leave out the agar. For 1 liter medium

  • 3 g NaCl
  • 17 g agar
  • 2.5 g peptone
  • 1 ml cholesterol (5 mg ml-1 in 95% EtOH)
  • 975 ml H2O

Autoclave, and then add the following sterile solution (autoclaved)

  • 1 ml 1 M CaCl2
  • 1 ml 1 M MgSO4
  • 25 ml 1 M potassium phosphate (pH 6) (to avoid precipitation, mix between addition of MgSO4 and potassium phosphate

Need to add streptomycin (300 ng ml-1) if plate is used for seeding bacterial food E. coli OP50-1. Typically pour 60 x 15 mm plate and store NGM plates in plastic boxes with covers at room temperature.

Seeding NGM plates with OP50

Day 1: grow an overnight culture of OP50 in LB medium. Pick a single colony from an LB plate using a sterile tip (stored at 4C). Place tip in a sterilized Erlmeyer flask with approx. 200ml of LB broth. Shake at 37C.

Day 2: Add 100ul of the OP50 culture to NGM plates and swirl to distribute. Let it dry overnight on the bench. Plates can be stored for 3 Weeks in the cold room.

Plate size OP50 Volume
3.5 cm 34 ul
6 cm 100 ul
10 cm 275 ul

Imaging plates

These are modified versions of the standard NGM plates . For 1 liter medium

  • 3 g NaCl
  • 17 g agarose (replace agar with agarose)
  • 2.5 g peptone
  • Do not add Cholesterol. It creates background
  • 975 ml H2O

Autoclave, and then add the following sterile solution (autoclaved)

  • 1 ml 1 M CaCl2
  • 1 ml 1 M MgSO4
  • 25 ml 1 M potassium phosphate (pH 6) (to avoid precipitation, mix between addition of MgSO4 and potassium phosphate

S-basal medium

For liquid culture of worms and use in microfluidics.

For 1 liter medium

  • 5.8 g NaCl
  • 50 ml 1 M potassium phosphate (pH 6)
  • 1 ml cholesterol (5 mg ml-1 in 95% EtOH)
  • 950 ml dH2O

Autoclave, and then add the following sterile solution (autoclaved)

  • 3 ml 1 M CaCl2
  • 3 ml 1 M MgSO4
  • 10 ml trace metals solution
  • 10 ml 1 M potassium citrate (pH 6.0)
  • 10 ml 100x Nystatin (antifungal agent, keep in freezer; we rarely add this).

500 ml trace metals solution

  • 0.346 g FeSO4.7H2O
  • 0.930 g Na2EDTA
  • 0.098 g MnCl2.4H2O
  • 0.144 g ZnSO4.7H2O
  • 0.012 g CuSO4.5H2O

Sterilize by autoclaving. Keep in dark (wrap in foil).

100 ml of 1 M potassium citrate

dissolve 21.02 g citric acid, monohydrate in 80 ml and adjust to pH 6.0 with solid KOH (approx 17g) before bringing up to volume.

Worm M9 buffer

  • 3 g KH2PO4
  • 6 g Na2HPO4
  • 5 g NaCl

Add H2O to 1 liter. Sterilize by autoclaving. After solution cools down, add

  • 1 ml autoclaved/sterile 1 M MgSO4.

Jun's note: you can also make 10X M9 solution by X10 of each components.

Worm lysis solution

This makes 100 ml 2x worm lysis solution. It is used for worm egg synchronization.

  • 50 ml ddH2O
  • 10 ml 10 M NaOH
  • 40 ml Clorox bleach

Make fresh and store at 4 °C up to one week.

Spot bleach protocol by Jun This protocol is to be used if you want to de-contaminate your worms when there are fungi or bacteria contaminations on the plate.
1) Add ~50ul of bleach solution to a seeded NGM plate. Avoid adding to the bacterial lawn.
2) Pick >5 gravid adults from contaminated plates and add the worms to the bleach solution on the plate.
3) Use the worm pick to press the worms a bit to help seperate the worms. Alternatively, you can also shake the plate gentally to help break-off.
4) leave overnight and the hatched L1 will crawl out.
5) Optional: transfer at least 4 L1s to a new plate just in case there are fungi survived (which occasionally happen when the original plates are heavily contaminated.

Canonical bleach protocol by Jun This protocol is to be used if you want to collect many synchronized L1s or to de-contaminate your worms when there are fungi or bacteria contaminations on the plate (this will give you many more embryos than the spot bleach).
Note: For behavioral assays, it is NOT recommended to use bleaching to obtain synchronous worms. Use egg laying method instead.\\. 1) Wash off 1 plate with many gravid worms with 1ml of M9.
2) Add them to a 1.5ml of Eppendorf tubes.

  • . Label the tubes well. The label might come off during the washes in the following steps.

3) Spin down at 1600g for 30~60sec.
4) Aspirate the supernatant and wash with 1ml of M9.

  • . It is ok to leave 50~100ul of liquid at the bottom. Otherwise you risk of losing worms.

5) Repeat step 4 for 2~3 times (until supernatant is clear).
6) Aspirate and add 1ml of bleaching solution.
7) Vortex or shaking in hand for 2min.
8) Spin down at 1600g for 1min.
9) Aspirate and add 1ml of bleaching solution.
10) Vortex or shaking in hand until you can see embryos under the microscope and that the worms are completely broken.

  • . No longer than 2min, otherwise you will kill the embryos.
  • . If you are just doing this for de-contamination, it is ok to have unbroken worms. If you are doing it for synchronization, note that the broken worms can be food for newly hatched L1 worms, thereby causing asynchronous population.

11) Spin down at 1600g for 1min.
12) Wash with 1ml of M9 at least three times.
13) Add 1ml of M9 and rotate at room temperature overnight.

  • . If you do not need to synchronize worms, you may just add the embryos from step 12 onto a new plate.

Bleaching solution (recipe for 100ml) by Jun

30 ml	Sodium hypochloride
15 ml	5M KOH 
55 ml	ddH2O

It always works the best when made fresh. However, it can be stored at 4⁰C for a week.

Freezing C. elegans using Liquid Freezing Solution

(from wormbook)

Equipment and Reagents

  • S Buffer [129 ml 0.05 M K2HPO4, 871 ml 0.05 M KH2PO4, 5.85 g NaCl]
  • S Buffer (see above) + 30% glycerin (v/v) (autoclave)
  • 1.8 ml cryotube vials
  1. Use one large, 2-3 medium, or 5-6 small NGM plates that have lots of freshly starved L1-L2 animals. Wash the plates with 0.6 ml S Buffer for each vial you will freeze. Collect liquid in a sterile test tube.
  2. Add an equal volume of S Buffer + 30% glycerin. Mix well.
  3. Aliquot 1.0 ml of mixture into 1.8 ml cryovials labelled with strain name and date.
  4. Pack the cryovials in a small styrofoam box with slots for holding microtubes or use a commercial styrofoam shipping box.
  5. Place the box in a −80°C freezer overnight (or for at least 12 hours).
  6. The next day transfer the vials to their permanent freezer locations. Thaw one vial as a tester to check how well the worms survived the freezing

Freezing C. elegans using Soft Agar Freezing Solution

Equipment and Reagents

  • Soft Agar Freezing Solution [0.58 g NaCl, 0.68 g KH2PO4, 30 g glycerol, 0.56 ml 1 M NaOH, 0.4 g agar, H2O to 100 ml (autoclave)]
  • 1.8 ml cryotube vials

Methods

  1. Melt Soft Agar Freezing Solution in autoclave or microwave and place in 50°C water bath for at least 15 minutes.
  2. Use one large, 2-3 medium, or 5-6 small NGM plates that have lots of freshly starved L1-L2 animals. Wash the plates with 0.6 ml S Buffer for each vial you will freeze. Collect liquid in a covered sterile test tube and place in ice for 15 minutes.
  3. Add an equal volume of Soft Agar Freezing Solution to the test tube. Mix well.
  4. Aliquot 1 ml of mixture into 1.8 ml cryovials labelled with strain name and date.
  5. Pack the cryovials in a small styrofoam box with slots for holding microtubes or use a commercial styrofoam shipping box.
  6. Place the box in a −80°C freezer overnight (or for at least 12 hours).
  7. The next day transfer the vials to their permanent freezer locations. Take a scoop of frozen mixture from one vial as a tester to check how well the worms survived the freezing

C. elegans growth speed

How to stage C. elegans

Determination of larval and adult stages (with Ingo) [Probably written by Florian Aeschimann]

Hallmarks of the different larval and adult stages of C.elegans visible with DIC under the Zeiss microscope (63x objective).

General remarks: Worms on the plates most of the time lie on their sides, the gonads/vulva are thus seen on one side (ventral), whereas the alae are seen in the middle of the worm surface (lateral).

Annuli are visible in all larval and adult stages.

Alae occur only on the cuticula of the L1 larvae and the adult worms (although the alae of the adult cuticula are already visible in late L4).

L1 Small gonad with only few gonad cells (only Z1-Z4 (picture C) in early L1, with their descendants in late L1) Alae on the worm surface

L2 Small gonad with an increased number of germ cells (picture D) No alae on the worm surface

L3 Larger gonad (extended gonad arms) with many germ cells (picture F) No vulva

Early L4 Gonad arms are reorientated (visible turn/bending is lab definition for early L4, otherwise still L3) Vulva is seen as a very small invagination

Mid L4 Gonad arms turn back about halfway Vulva is seen with a Christmas tree structure (picture G)

Late L4 Gonad arms turn back all the way Vulva is seen with a Christmas tree structure or further developed, but with a cuticula covering the vulva Alae structures are already visible from the adult cuticula below the L4 cuticula

Young adult Gonad arms turn back all the way and overlay each other Vulva is developed, without a covering cuticula Alae on worm surface No embryos visible

Adult Gonad arms turn back all the way and overlay each other Vulva is open to the outside (picture H) Alae on worm surface Embryos visible (about 4 hours after L4-adult molt?)

To precisely stage L3 and L4 worms, see below for vulva morphology comparison. Please note the timing in the following picture refers to the growth rate at 22 degrees. PMID: 24945623

wiki/protocols.txt · Last modified: 2020/03/26 11:32 by jliu

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