Cagan Laboratory

Collect Bugs:

1.Pour 2ml overnight culture into 1.5ml centrifuge tube

2.Spin 30 secs; pour off supernatant

3.Resuspend in remaining broth by vortexing

Extracting Plasmid:

4.Make fresh STET/lysozyme solution
          •  STET: 8% sucrose + 50mM Tris8 + 50mM EDTA + 5% TritonX-100
          •  can store at room temperature
          •  Add fresh lysozyme powder to 1mg/ml final concentration

5.Vortex in 400µl STET/lysozyme solution
          •  remember: have large vortex holder

6.Boil 1’

7.Spin 10’ @ R.T.

8.Remove pellet with toothpick or yellow tip

Cleaning up Plasmid:

9.Add 400µl chilled isopropanol; vortex.

10.Spin 15’ @ 4°C.

11.Wash pellet with 70% EtoH.

12.Lyophilize.

13.Resuspend in 50-100µl TE8. Use 1µl per miniprep.

2.Resuspend in 50 µl of lysis buffer
          •  10mM Tris-8 + 1mM EDTA + 15% sucrose + 2mg/ml lysozyme + 0.2 mg/ml RNase + 0.1 mg/ml BSA
          •  store at -20°C

3.Boil 1 min.

4.Chill 1 min.

5.Spin 15-20 min @ R.T.
          •  Can leave pellet @ bottom and draw from supernatant. Remove pellet for longer-term storage.

6.Use 3µl-5µl for digestion.

1.Fix tissue in PLP or PEM for 30’.

2.Wash 1×10’ in PBS; then 2×10’ in 0.1% saponin/PBS
          Never use Na Azide. Can use 0.01% thimerosol

3.1° Ab + 10% goat serum in 0.1% saponin overnight

4.Wash 3 x 10’ in saponin

5. If using (eg) HRP-conj 2° Ab:
          1.  2° Ab + 10% goat serum in 0.1% saponin 1-2 hrs.
     If using vectastain:
          1.  2° biotin-Ab + 10% serum in saponin 1-2 hr
          2.  Wash 3×10’ in saponin
          3.  “A” (1:100) + “B” (1:100)  + 10% serum in saponin 1-2 hr

6. Wash 2×10’ in saponin, then 2×10’ in PBS.
Wash tissue thoroughly before refixing

7. Fix 15’ in 2% glutaraldehyde/PBS; wash 3 x 10’ in PBS

8. Develop with 0.5 mg/ml DAB in PBS
          Can add 5µl of 8% NiCl2 solution to 1 ml DAB to enhance contrast
          Stop reaction by transferring discs to PBS with 0.01% Na Azide

9. Wash 2×10’ in PBS; mount in Aquamount

High penetration:

1. Fix tissue in 4% paraformaldehyde for 20 min.

2. Wash 1×10’ in PBS; then 2×10’ in 0.3% Triton-X100/PBS
          Never use Na Azide. Can use 0.01% thimerosol
          Note that Triton-X100 is the same as NP40

3. 1° Ab + 10% goat serum in 0.3% Triton-X100 for  2 hrs-overnight; shaking

4. Wash 2x 10’ in Triton-X100

5. 2° Ab + 10% goat serum in 0.3% Triton-X100 2 hrs; shaking

6. Wash 2×10’ in 0.3% Triton-X100, then 2×10’ in PBS.
          Wash tissue thoroughly before refixing

7. Fix 5’-10’ in 2% glutaraldehyde/PBS 

8. Develop with 0.5 mg/ml DAB in PBS
          Can add 5µl of 8% NiCl2 solution to 1 ml DAB to enhance contrast
          Stop reaction by transferring discs to PBS with 0.01% Na Azide

9. Wash 2×10’ in PBS; mount in Aquamount

1.dissect in PBS

2.fix in PLP for 30’

3.Wash 3X10’ in PBS

4.fix 30’ in 2% glutataraldehyde (from freezer) in PBS

5.wash 3X 10’ in PBS
          post-fix in OsO4/K3Fe(CN)6 for 60’
          add equal volumes of OsO4 and K3Fe(CN)6 from tubes in freezer container marked “Microscopy”.

6.Wash 3X10’ in PBS

7.Dehydrate in EtOH series: 30%, 50%,70%,80%,95%, and 3X100%
!!!!!!!!!No cheating!!!!!!!!!!!!!! This is critical for EM!!!!!!!

8.3X10’ in propylene oxide

9.overnight in 1:1 propylene oxide:EM resin
          use gloves

10.5 hrs in 100% resin; imbed