1. ß-galactosidase and hPLAP staining - solutions
2. Genotyping by lacZ staining
3. lacZ wholemount staining
4. hPLAP wholemount staining
5. lacZ and hPLAP slide staining
6. Genotyping by Southern
7. in situHybridization to Whole Mount Embryos
8. in situHybridization to tissue sections with DIG-labelled probes
9. DNA for Microinjection

Solutions

0.1 M NaPO4 buffer: (for 500 ml)

11.5 ml 1 M monobasic NaPO4
38.5 ml 1 M dibasic NaPO4
450 ml water

lacZ fix: (for 50 ml)

0.4 ml 25% glutaraldehyde
1.0 ml 250 mM EGTA pH 7.3
5.0 ml 1 M MgCl2
43.5 ml 100 mM sodium phosphate, pH 7.3 or PBS

Note: if doing a wholemount hAP stain, a mixture of 0.2% glutaraldehyde in PBS + 0.02% NP-40 + 0.01% NaDC may improve penetration of stain

lacZ wash buffer: (for 500 ml)

1. ml 1 M MgCl2
5.0 ml 1% sodium deoxycholate (NaDC; make up in water; store in fridge)
5.0 ml 2% Nonidet-P40 (make up in water; store in fridge)
489 ml 100 mM sodium phosphate (pH 7.3)

lacZ stain: (for 75 ml)

72.0 ml wash buffer
3.0 ml 25 mg/ml X-gal (dissolved in DMSO)
0.159 g K-ferrOcyanide
0.123 g K-ferrIcyanide

AP wash buffer: (for 100 ml)

10 ml 1 M Tris-HCl, pH9.5
2 ml 5 M NaCl
1 ml 1 M MgCl2
87 ml water

NBT/BCIP stain: (for 20 ml)

2 ml 1 M Tris-HCl, pH9.5
0.4 ml 5 M NaCl
1 ml 1 M MgCl2
200 µl 1% sodium deoxycholate (final 0.01%)
200 µl 2% NP-40 (final 0.02%)
70 µl NBT (to get 337 µg/ml)
70 µl BCIP (175 µg/ml)
16 ml water

top of page

Genotyping by lacZ expression

Earpunch mice and place biopsies into PBS in a 96-well plate
Fix 5 to 30 minutes (usually 15 minutes) in 0.2% glutaraldehyde in PBS (can just add equal volume of 0.4% glutaraldehyde in PBS)
Wash 3 times for 5 minutes in PBS
Stain with X-gal stain at 37 C (or at room temp) protected from light
Staining is usually visible after 5 minutes if fresh stain is used

top of page

lacZ wholemount staining

Rinse embryos/tissues in 100 mM sodium phosphate (pH 7.3) or PBS

Fix:
- small embryos (<E9.5) in lacZ fix for 15 to 30 minutes on ice, with gentle shaking,
- large embryos in 2% paraformaldehyde and 0.2% glutaraldehyde in PBS for 30 minutes on ice with shaking, then bisect and fix for an additional 30 to 60 minutes on ice in lacZ fix
- large tissues in lacZ fix for 4 hours on ice with shaking, bisecting the tissue after the first hour to allow penetrance of the fix solution.

Wash 3 times for 15 to 30 minutes in lacZ wash buffer.

Stain in lacZ stain solution at 37oC or room temperature for 30 minutes to overnight, with shaking and protected from light.

Wash 3 times for 10 minutes in PBS and store in lacZ wash buffer at 4 C.

top of page

hPLAP wholemount staining

Pre-heat a waterbath to 70 C and place a bottle/tube with the required amount of PBS in it

Fix: For Alkaline Phosphatase staining, embryos and tissues are fixed as for the lacZ wholemount protocol above. For better penetration of the Alkaline Phosphatase substrates, 0.02% NP-40 and 0.01% sodium deoxycholate can be added to the fix solution.

Rinse in PBS

Heat inactivate endogenous alkaline phosphatases by incubation in PBS at 70 to 75oC for 30 minutes.

Rinse samples in PBS

Wash in AP buffer for 10 minutes

Stain in NBT/BCIP stain, protected from light, at room temperature for 10 to 20 minutes or at 4 C for 0.5 to 36 hours.

Wash extensively in PBS containing 0.1% Tween-20 and 2 mM MgCl2.

top of page

lacZ and hPLAP slide staining

Slide preparation:

Dissect samples into cold PBS

Fix samples as for wholemounts. Petri dishes or 6-well plates work well for containers

Wash samples 3 x 20’ in PBS

Cryoprotect in 15% sucrose/PBS for 1 hour at 4oC

30% sucrose/PBS overnight at 4oC.

Place in Tissue-Tek OCT (Sakura) at 4oC for several hours (at least 1 hour)

Embed in OCT:
put OCT into a plastic mold (write a label on the mold first)
place the tissue/embryo into the mold and orient it
place the mold onto dry ice (best to have some dry ice in a plastic tray) and hold it level, being careful to maintain the orientation of the embryo

When all samples have been prepared, wrap them in saran wrap, place in a labeled box and store at -80

Cryosection blocks at 7 to 10 µm (give the blocks about 30 minutes in cryostat chamber to equilibrate to —20). Place the sections onto poly-lysine coated slides (Fisher Scientific)

Dry slides for 1 to 4 hours at room temperature

Store at -20oC in a slide box, with a little dessicant (Dry-rite, should be blue) and taped shut

To stain:

Take slides out of —20 and equilibrate to room temp before opening the box

If doing hAP stain, place a large staining jar with PBS in a waterbath and pre-heat to 70 C

Place slides to be stained into staining jars (hold 15 slides and take 75 ml). Leftover slides can be placed back into —20 with dessicant.

lacZ stain

Fix slides in cold PBS containing 0.2% glutaraldehyde for 10 minutes

Wash 3 times for 5 minutes in lacZ wash buffer

Stain in lacZ stain solution for 4 to 6 hours at 37oC, protected from light (wrap foil around the staining jar and place in a TC incubator).

When the staining is complete, rinse slides 3 x 5’ in PBS

If only doing lacZ stain, at this point dehydrate the slides and mount coverslips (see below)

hAP stain

For hAP staining alone, start with fix, then 3 times 5’ washes in PBS, then proceed as follows:

Inactivate endogenous alkaline phosphatase by incubating slides in PBS at 70 to 75oC for 30 minutes, in the preheated staining jar.

When finished, let slides cool at rm temp for 1 to 2 minutes.

Rinse with PBS

Wash in AP buffer for 10 minutes

Shake excess liquid from slides and place on the rack in a Tupperware "pizza box"

Overlay with NBT/BCIP stain (requires 0.5 to 1 ml per slide). Put the cover on the box and cover with foil to protect from light

Stain for 10 to 30 minutes at room temperature

Wash the slides 3 times for 10’ in PBS/2mM MgCl2

Dehydrate through an ethanol series:

5’ PBS
5’ 70% etOH/PBS
5’ 90% etOH/PBS
5’ 100% etOH
5’ 1:1 etOH/xylene (use xylene in the fume hood)
5’ xylene
5’ xylene

Mount with coverslips

Normally counterstain is not a good idea, but if desired insert the following two steps before the dehydration steps above:

5’ PBS
30’’ to 5’ Nuclear Fast Red (5% AlSO4, 0.1% Fast Red)

Length of stain depends on freshness of NFR

top of page

The Cre coding sequence can be used as a radioactive probe for Cre transgenics or PCR primers can be used to amplify the probe. Primer 1 is ATG TCC AAT TTA CTG ACC G; Primer 2 is CGC CGC ATA ACC AGT GAA AC.

top of page

in situHybridization to Wholemount Embryos

A. Preparation of Single-Stranded RNA Probes

1. Mix at room temperature (following this order):

Sterile distilled Water 10.0 ul
5 X Transcription buffer 4.0 ul
0.1 M DTT 2.0 ul
Nucleotide mix 2.0 ul
10.0 mM GTP
10.0 mM ATP
10.0 mM CTP
6.5 mM UTP
3.5 mM digoxigenin-UTP
Linearized plasmid (1mg/ml) 1.0 ul
RNasin (100 u/ml) 0.5 ul
SP6,T7, OR T3 RNA polymerase (10U/ml) 1.0 ul

2. Incubate 37 C for 2 Hours

3. Add 100 ul TE

4. Remove 5 ul for gel analysis. To the 5 ul aliquot, add 1 ul RNA loading dye (50% formamide plus BPB/XC) and load on a 1% agarose TBE mini-gel. The regular DNA molecular weight marker should be used. Run the gel at 100 V for 30’ to 60’.
You should see the DNA template band and the smaller RNA band. The RNA band should be 5 to 10 times more intense than the DNA

5. Precipitate remainder with 10 ul 4M LiCl + 300 ul EtOH

Leave at -20 C for 30 min.

6. Centrifuge for 10 min.

7. Wash pellet with 70% EtOH and BRIEFLY air dry.

8. Redissolve in 100 ul TE (to 0.1 mg/ml)

9. Store the DIG probe at -20 C. Label the tube with the expiry date of the DIG mix used in case the probe outlasts the DIG mix.

B. Preparation of embryos

1. Prepare 4% paraformaldehyde (PFA) as follows: For 100 ml, weigh out 4 g of PFA (caution: do not breathe dust). In the fume hood, put the PFA in a 500 ml erlenmeyer flask with a clean stirring bar. Add 86 ml ddH2O. Heat with stirring, while monitoring the temperature, until the solution reaches 70 C. Remove from heat. Add 2 to 3 drops of 5 M NaOH and swirl to mix. The solution should go clear. Note: DO NOT ALLOW THE PFA TO BOIL.
When the solution cools, add 10 ml of 10 x DEPC-PBS and make the final volume up to 100 ml. Store at -20 in 10 ml aliquots.
When ready to dissect embryos, thaw a tube of 4% PFA and place on ice. Do not re-freeze any unused solution - dispose of it.

2. Sacrifice the pregnant mouse and remove the uterus to a petri dish of cold PBS. For E7 to E9, dissect away excess connective tissue and fat, then cut the length of uterus into individual embryos. Leave the dish on ice, and one at a time, remove the embryos to a fresh petri dish containing PBS under the dissecting microscope. Dissect each embryo out of the decidua and membranes, immediately place the embryo in the PFA using a cut-off blue tip/pipetteman, then dissect the next embryo.

3. Fix the embryos for 4 hours to overnight at 4 C.

4. Wash and dehydrate the embryos as follows: (PBT is PBS / 0.1% Tween)

PBT 5 min 4 C
PBT 5 min 4 C
25% MeOH/PBT 5 min
50% MeOH/PBT 5 min
75% MeOH/PBT 5 min
100% MeOH 5 min
100% MeOH store at -20 C

C. Hybridization - Day 1

Carry out all steps in 1.5 ml O-ring tubes or 15 ml snap-cap tubes WITH GENTLE SHAKING, unless otherwise indicated.
Initially take out the embryos you require from the stock at -20 in MeOH and sort into tubes, one tube per RNA probe. Alternatively, you can process them in one tube per stage (ie d7, d8, d9) and re-sort them before pre-hybridization.
For the 1.5 ml tubes, use 1 ml of each solution per tube.
Do not put, for example, E7 with E9 embryos, as they will require different lengths of Prot K treatment.

1. 75% MeOH in PBT 5 min

2. 50% MeOH in PBT 5 min

3. 25% MeOH in PBT 5 min

4. PBT 5 min.

5. PBT 5 min.

6. 6% (v/v) H202 in PBT 1 hour (use H202 that has been open < 1 month)

7. PBT 5 min

8. PBT 5 min


9. PBT 5 min

10. 20 ug/ml proteinase K in PBT 2 to 15 min (for 7 to 9 dpc embryos)

11. 2 mg/mL glycine in PBT (fresh) 5 min.

12. PBT 5 min.

13. PBT 5 min.

14. 0.2% glutaraldehyde/4% PFA in PBT 20 min
(Add 8 ul 25% glutaraldehyde and 5 ul 20% Tween per 1 ml 4% PFA)

15. PBT 5 min.

16. PBT 5 min.

17. pre-hybridization sol’n 1 hour at 70 C (use temp block on side or Hybaid oven)
(50% formamide, 5 X SSC,pH 5, 50 mg/mL yeast RNA, 1%SDS, 50 mg/ml heparin)

18. pre-hybridization solution + 10 ul probe overnight at 70 C

D. Hybridization - Day 2

19. Solution 1 30 min. 70 C
(Solution 1: 50% formamide, 5X SSC, pH 4.5, 1% SDS)

20. Solution 1 30 min. 70 C

21. 1:1 solution 1:solution 2 10min 70 C

22. Solution 2 5 min rm temp
(Solution 2: 0.5 M NaCl, 10mM Tris-HCl pH 7.5, 0.1% Tween 20)

23. Solution 2 5 min

24. Solution 2 5 min

25. Solution 3 5 min
(Solution 3: 50% formamide, 2 X SSC,pH 4.5)

26. Solution 3 30 min 65 C (start on step 32 in parallel now)

27. Solution 3 30 min 65 C

28. TBST 5 min.
(10X TBST (100 ml): 8 g NaCl, 0.2 g KCl, 25 ml 1 M Tris-HCl pH 7.5, 10 ml Tween-20. Autoclave. Dilute to 1x and add levamisole to 2mM (0.48 mg/ml) on day of use)

29. TBST 5 min.

30. TBST 5 min.

31. Preblock embryos by incubating with 10% sheep serum in PBT for 2 - 3 hours.
(Incubate serum 30 min at 70 C once before use)

32. Preabsorb antibody with embryo powder as follows:
- 3 mg embryo powder in 0.5 mL TBST, 30 min @ 700C with shaking (use shaking heater block)
- place on ice and allow to cool for a few minutes
- add 5 ml sheep serum, 1 ml anti-DIG antibody
- rock gently for 1 hour at 40C
- spin 5’
- Remove supernatant to fresh tube and dilute to 2 ml with 1% serum/TBST

33. Remove 10% serum/TBST from embryos

34. Replace with preabsorbed antibody

35. Rock gently O/N at 4 C

E. Hybridization - Day 3

36. TBST 5 min

37. TBST 5 min

49. TBST 5 min

50. TBST 1 Hr

51. TBST 1 Hr

52. TBST 1 Hr

53. TBST 1 Hr

54. TBST O/N

F. Hybridization - Day 4

56. NTMT 10 min.
(NTMT: 100 mM NaCl, 100 mM Tris-HCl pH 9.5, 50 mM MgCl2, 0.1% Tween-20, 2mM levamisole, make from stocks on day of use)

57. NTMT 10 min.

58. NTMT 10 min.

59. Transfer the embryos to a 6-well TC dish.

60. Incubate embryos in NTMT + 3.5 ml NBT/ml + 3.5 ml BCIP/ml (you will need 3 ml/well)

61. Incubate w/ gentle shaking 20 min. (in the dark)

62. Leave 1Hr- 0/N in the dark. Signal usually is optimal at 4 to 6 hours

63. Wash PBT >15 min.

64. Wash PBT >15 min.

65. Wash 50% glycerol/PBT >15 min.

66. Sore in 80 % glycerol/PBT > 15 min. 40C

top of page

in situHybridization to tissue sections with DIG-labelled probes

A. Preparation of Embryos

1. Dissect embryos in PBS as described for whole mount in situs

2. Transfer embryos to 4% paraformaldehyde in PBS. Leave at 4 C 4 hours to overnight.

B. Embedding

3. Wash PBS 1 Hr.

4. Wash .86% NaCl (0.15 M) 1 Hr.

5. Wash 70% Ethanol 1 Hr.

(embryos can be stored in 70% EtOH until taken to pathology)
6. Take to Pathology, Histology Section in E 441 for them to be embedded in paraffin. Fill out a requisition, including a description of the orientation you wish the embryos to have in the block.

7. Pick up wax blocks the next day

C. Sectioning

Preheat histology water bath to 45 - 50 C. Chill tissue block at 4 C for easier sectioning.

8. Cut 7 to 20 mm ribbons (normally 7 mm is used, but for DIG in situs, the thicker sections provide better sensitivity)

9. Place ribbons on bath of dH20 at 500C until creases disappear

10. Collect on subbed slides

11. Dry sections onto slides 37C O/N

12. Store slides in slide box at 4 C.

DAY 1: Prehybridization and Hybridization of Tissue Sections

Prehybridization of slides is carried out in glass jars with 250 ml of each solution. See end of protocol for wash and treatment of glass jars. Set the jars out in rows, with a line of tape underneath. Write the solution and time on the tape under each jar.

1. Histoclear or Xylene 10’

2. Histoclear or Xylene 10’

3. 100% EtOH 2’

4. 100% EtOH 2’

5. 100% EtOH 2’

6. 95% EtOH 2’

7. 90% EtOH 2’

8. 80% EtOH 2’

9. 70% EtOH 2’

10. 50% EtOH 2’

11. 30 % EtOH 2’

12. Saline 5’ (.86% (150 mM) NaCl)

13. PBS 5’

14. 6% H202 /PBS 30’ (use H202 that has been open for less than one month)

15. PBS 5’

16.PBS 5’

17.4%(w/v) paraformaldehyde in PBS 20’ make fresh

18.PBS 5’

19. PBS 5’

20. 20mg/ml proteinase K from 10mg/ml stock, diluted in 50 mM Tris-HCl, 5mM EDTA, pH8.0 1’ CAUTION: Not longer!!!!

21. PBS 5’

22.4%(w/v) paraformaldehyde in PBS 20’

23.PBS 5’

24.Acetylation mix 10’
250 ml 0.1M triethanol amine+ 0.625 ml acetic anyhydride make fresh

25. PBS 5’

26.Saline 5’

27. 30 % EtOH 2’ (re-use the ethanols from dehydration EXCEPT the 100%)

28. 50% EtOH 2’

29. 70% EtOH 2’

30. 80% EtOH 2’

31. 90% EtOH 2’

32. 95% EtOH 2’

33. 100% EtOH 2’

34. Air Dry

Slides are now ready for the probe.

Make up sufficient volume of Hybridization mix. You will need 100 ul/slideHybridization mix:

Final Stock Add/ml
50 % Formamide 100% 500 ul
5X SSC,pH 5 (Adj.with citric acid) 20X 250 ul
50 ug/ml yeast tRNA 10 mg/mL 5 ul
1 % SDS 10 % 100 ul
50 ug/ml Heparin 10 mg/mL 5 ul
H20 140 ul

-Dilute probe in hybridization mix at 5:95 (probe:hyb)

35. Put 100 ml of probe/hybridization mix per slide.

36. Add cover slip, make sure probe/hyb. mix covers all tissues.

37. Incubate in a Tupperware pizza-box O/N at 650C. Put ~100 ml of 50% formamide/5XSSC in bottom to maintain humidity in the box.

DAY 2: Washes and Antibody binding

38. Place slides in rack in 250 ml 5 X SSC 650C until coverslips fall off.

Carry out the following steps in the small slide jars. Each holds 80 ml.

39. Solution 1 65 C 30’
(50% formamide, 5X SSC pH 5.0, 1% SDS)

40. Solution 1 65 C 30’

41. Solution 2 Rm.Temp. 10’
(Solution 2: 0.5 M NaCl, 10mM Tris-HCl pH 7.5, 0.1% Tween 20)

42. Solution 2 Rm.Temp. 10’

43. Solution 2 Rm.Temp. 10’

44. Solution 2 37 C 45’

45. Solution 2 Rm.Temp. 5’

46. Solution 3 65 C 30’
(Solution 3: 50% formamide, 2 X SSC pH 5.0)
**Start preabsorption of antidigoxigenin antibody. See below

47. Solution 3 65 C 30’

48. PBT Rm temp 10’
(PBT: PBS, 0.1 % (v/v) Tween 20)

49. PBT 10’

50. 10% sheep serum in PBT 2-3 Hrs

51. PBT rinse quickly

52. Overylay sections w/ preabsorbed antibody and cover with coverslips. Place in a tupperware pizza box with some water on the bottom to keep the moisture.

53. Incubate in moist chamber O/N 40C

DAY 3: Washes and signal detection

54. PBT 5’

55. PBT 5’

56. PBT 5’

57. PBT 30’

58. PBT 30’

59. PBT 30’

60. NTMT 5’
(NTMT: 100 mM NaCl, 100 mM Tris-HCl pH 9.5, 50 mM MgCl2, 0.1% Tween 20, 2mM levamisole)

61. NTMT 5’

62. NTMT 5’

63. NTMT containing 3.5ul NBT, 3.5 ul BCIP per ml
NBT 100mg/ml stock in dimethylformamide
BCIP 50 mg/ml stock in dimethylformamide

64. Incubate in dark 5-6 hrs

Colour reaction can be monitored by briefly looking at a slide under the dissecting microscope using illuminated light (ie from above). DO put saran wrap over the base of the microscope first, to avoid staining it!

65. When sufficient colour has developed, rinse slides in PBT, then in ddH2O

67.Air-dry slides overnight in fresh, dry slide jars

68. Place 5 or 6 drops of Cytoseal on the slide and cover with cover slip (avoid air bubbles)

Preabsorption of Antibody

1. Weigh 3 mg embryo powder into microcentrifuge tube

2. Add 0.5 ml PBT

3. Heat 70 C 30’

4. Cool on ice

5. Add 5 ul sheep serum (previously heat-treated at 70 C for 30 minutes)

6. Add 1 ul anti-digoxigenin antibody

7. Shake gently 4 C 2-3 Hrs.

8. Spin in centrifuge 5’

9. Dilute supernatant to 2 ml with 1% serum in PBT

Embryo Powder

1. Dissect 12.5 to 14.5 day embryos in minimum amount of ice-cold PBS

2. Homogenize 2 X 30 seconds

3. Add 4 volumes ice-cold acetone to homogenate (the acetone should be from a fresh bottle or maintained anhydrous with molecular sieves) and mix by inversion

4. Incubate on ice for 30 min.

5. Centrifuge 10,000 g 10 min.

6. Remove supernatant. Re-suspend pellet in ice-cold acetone

9. Centrifuge 10,000 g 10 min.

10. Remove supernatant and spread pellet out in a RNase-free glass dish or petri dish. If necessary, grind it to a fine powder with an Rnase-free metal spatula

12. Air-dry powder overnight at room temperature with the cover on the dish

13. Store at 4 C

Glassware for in situs

Pour the used solutions from the glass jars down the sink, except Xylene, which goes in the special waste container in the fume hood. Do not used soap to clean the jars. Rinse the jars about 7 times with tap water, then 2 to 3 times with distilled water, then with ddwater. Set the rinsed jars on a sheet of banchcoat to drain.

The jars are then baked at 360 F/180 C overnight. When baking is finished, turn the oven off and allow the jars to cool before removing them - if you don't they will crack.

In addition, a stock of baked glass graduated cylinders and bottles should be prepared. Use foil to cover tops.

RNase-Free Stock Solutions

Make the following solutions from standard stocks and DEPC treat:

Sterile distilled water
4 M LiCl
10 X PBS
5 X SSC, pH 4.5
5 M NaCl
0.5 EDTA
1M MgCl2

Make the following solutions from RNase-free stocks in a baked glass bottle or RNase free plastic container and DO NOT DEPC-treat:

1 M Tris pH 7.5
TE
EtOH
70% EtOH
4% paraformaldeheyde
5 M NaOH
20% Tween
10% SDS
50 µg/ ml yeast RNA
50 µg/ ml heparin
RNA loading dye
10 X TBST
Embryo Powder

*Although the above items must not be DEPC treated, they can be made from DEPC-treated water

Supplies:

Nucleotide mix Boehringer Mannheim 1 277 073
Rnasin Promega N251A
SP6, T7, or T3 RNA Promega (all 3) P108 B, P207B, P208C
Levamisole Sigma L9756
Sheep serum Sigma S2263
anti-DIG antibody Boeringher 1 093-274
NBT Boeringher 1 383 213
BCIP Boeringher 1 383 221
Glycerol MDC 10816
Xylene Fisher X5B-500
0.1 M triethanol amine Sigma T9534
Acetic anyhydride Sigma A6404
Slides polyprep slides Sigma P0425
Cover slip, glass 22 X 50 VWR 48393 059
Hybri-slips Sigma Z37,027-4
Heparin Sigma H3149
Tween-20 BDH R06435
H202 Sigma H1009
DEPC Sigma D5758
50% formamide Sigma 18,590-6
1.5 ml 0-ring Diamed TEC 152SC-N
Proteinaise K Boeringher 745 723
Glycine Sigma G6388
Glutaraldehyde BDH B28682
Tupperware pizza-box Dollar store
Alkaline Phosphatase Boeringher Mannheim 713 023

top of page

NOTE: For microinjection at the SWCHSC transgenic facility, we require a minimum of 20uL of DNA that is at least a concentration of 25ng/uL.

This protocol uses the CONCERT Matrix Gel Extraction System from Gibco and Elutip column from Mandel Scientific. It is a rapid and clean way of making injection fragments, but fragments of over 10kb may be sheared. It is important to use a microfuge that accelerates quickly (full speed in under 10sec) to limit the shearing forces.

Gel purify from Gibco maxiprep DNA:

1. Digest about 10-20ug plasmid. Separate from the vector backbone on a 0.7% agarose gel (with 5uL/100mL gel of 10mg/mL Ethidium bromide) made in 1X TAE. Run at about 70V.

2. Cut out the band under UV light (keep UV exposure short and at 300nm wavelength). Turn gel slice on its side and trim off excess that does not contain DNA.

3. Follow the Gibco CONCERT Matrix Gel Extraction System protocol to extract the microinjection DNA fragment from the gel.
a. Place up to 350mg gel slice into 1.5mL polypropylene tube. Divide gel slices exceeding 350mg among additional tubes. Add 30uL Gel Solubilization Buffer (L1) for every 10mg gel.
b. Vortex the Silica Resin until it is thoroughly suspended. Add 1uL of Resin for every 10mg gel. Vortex and incubate at 50C for at least 15min. Mix every 3min to ensure gel dissolution. After gel slice appears dissolved, incubate for 5min longer.
c. Centrifuge at least 12000g for 30sec. Carefully and thoroughly remove supernatant with a pipette and discard.
d. Add 30uL L1 buffer for every 10mg gel. Suspend resin by vortexing or by flicking the tube. Centrifuge at least 12000g for 30sec. Carefully and thoroughly remove supernatant with a pipette and discard. This is the high-salt wash.
e. Add 30uL Wash Buffer (L2) containing ethanol for every 10mg gel. Suspend resin by vortexing or by flicking the tube. Centrifuge at at least 12000g for 30sec. Carefully and thoroughly remove supernatant with a pipette and discard. This is the low-salt wash.
f. Repeat the low-salt wash.
g. Air dry the silica resin to remove residual ethanol from the wash buffer. Do not overdry.
h. Resuspend the resin in 20uL sterile TE. Incubate at 50C for 5min to elute DNA. Mix once during the incubation.

Clean up the DNA on an Elutip column from Mandel Scientific:
1. Dilute the DNA with H2O to total of 960 ul and add 40 ul 5M NaCl.
2. Prepare the colunn by running through 3 ml of High salt buffer, and then 3 ml of Low salt buffer, by attaching the column to 3 ml syringes loaded with these buffers (see kit for recipes).
3. Run your DNA sample through the column at about 1 drip per second using a 3 ml syringe. The DNA binds to the column.
4. Wash the bound DNA using 3 ml of Low salt buffer (1 drip per second).
5. Elute the DNA into a microfuge tube using 450 ul of High Salt buffer in a 1 ml syringe (1 drip per second).
6. Add 900 ul 100% ethanol. Mix and precipitate overnight at -70 C.
7. Spin down DNA 15 mins in microfuge in cold room and resuspend in 25 ul injection buffer (10 mM Tris pH 7.5, 0.2 mM EDTA pH 8.0) prepared in high quality Milli-Q H2O. Run 2 ul on a minigel with molecular weight markers of known concentration to get an estimate on the amount of DNA. Make sure that the DNA is intact. Estimate the DNA concentration of your sample by comparison to the marker DNA.

The DNA will be diluted to 2 to 4 ng/ul for injection, but must be supplied at a minimum concentration of 25 ng/ul.

top of page