Table of Contents

Protocols

See also: How-to
See also: Commercial Protocols (GitHub)
See also: Product Information Sheets
See also: JGI Protocols
See also: NGS Protocols



General Reagents



How to make a solution:

1. Calculate the quantity of solute needed for your desired concentration and volume using one of the following formulas:

Solute needed for desired molar solution: MW of solute (g/mol) x Molarity (mol/L) x Volume (L) = grams of solute
Solute needed for desired percent solution: 1% solution = 1g solute/100mL total volume
Volume needed for desired percent solution: 1% solution = 1mL reagent/100mL total volume

2. Fill beaker with no more than half the final volume of the proper solvent. The solvent is molecular grade water, unless otherwise noted.

3. Add stir bar to beaker, place on stir plate and begin stirring.

4. Gradually add the solute to the solvent. Wait until the solute you added has dissolved, then add more solute. Repeat until solute is dissolved.

5. Rinse weigh boat with molecular grade water into the beaker containing your solution.

6. Calibrate pH meter and adjust the pH (See "How-to" for instructions on using the pH meter.)

7. Transfer solution to graduated cylinder and top off solution to desired volume.

8. Transfer solution from graduated cylinder to proper storage container.

9. Label container with solution name (not just chemical formula!), concentration, pH, date and your initials.

9. Autoclave solution (See "How-to" for instructions on autoclaving solutions), unless otherwise noted.

10/27/2011 SJW




TE Buffer

10mM Tris-HCl
1mM EDTA

Adjust pH to 8.0



10x DNA Loading Buffer

Dissolve in TE:

0.42% Bromophenol Blue
0.42% Xylene cyanol
25% Ficoll

Ficoll takes a LONG time to go into solution. Extensive vortexing will be required.

Coomassie Blue Stain (1L)

450mL methanol
100mL acetic acid
400mL water
2.5g Coomassie brilliant blue
D/W up to 1L

10x TBE Buffer

108 g Tris Base
55 g Acid
9.3 g Na2 EDTA×2H2O or 40 ml 0.5M EDTA pH 8.0
D/W up to 1 L
Needs to be pH’ed

10x Modified TAE Buffer

48.4 g/L Tris Base
11.4 ml/L Glacial Acetic Acid
2 ml/L 0.5 M EDTA

NOTE: Cannot be autoclaved; acetic acid will evaporate.
NOTE: In modified TAE buffer, the 1x working solution will contain 0.1mM EDTA instead of the standard 1mM EDTA.

20x SSC

Dissolve 175.3 g NaCl + 88.2 g Sodium Citrate in 800 ml DEPC H2O
Adjust pH to 7.0 with HCl or NaOH
Adjust the volume to 1 L with DEPC H2O
Filter
Do not need to autoclave
(For 500 ml use 87.65 g NaCl + 44.1 g Sodium Citrate)
(For 1.5 L use 262.95 g NaCl + 132.3 g Sodium Citrate)

LB Broth

10 g NaCl
10 g Bactotryptone
5 g Yeast Extract
Mix on stir plate up to 800 ml with D/W
pH to 7.0
Add D/W to 1 L
Autoclave

LB Plates

10 g NaCl
10 g Bactotryptone
5 g Yeast Extract
Bring to 800 ml with water and stir on stir plate
Adjust pH to 7.0
Add 20 g Bacto Agar
Autoclave
Cool at 50C in waterbath
Once at 50C add antibiotics if necessary – for Kan use 1 ml Kan per 1 L solution
Stir
Pour plates
Flame bubbles off

NZY Broth

5 g NaCl
2 g MgSO4 × 7 H2O (or 1 g MgSO4 Anhydrous
5 g Yeast Extract
10 g N2 Amine (Casein Hydrolysate)
Add water up to 800 ml
Adjust to pH 7.5 with 1 N NaOH (about 1-2 ml)
Add water up to 1 L
Autoclave or filter sterilize

NZY Plates

Add 15 g Difco Agar / liter of NZY Broth
Autoclave
Use about 80 ml per large plate; about 35-40 ml per small plate
Cool to 50°C to pour or to add antibiotics
For NZY + amp Plates:
50 mg/ml ® 1 ml/L = 50 ug/ml
For NZY + kan Plates:
50 mg/ml ® 1 ml/L = 50 ug/ml

Top Agar

Make 1 L NZY broth and add only 7 ml agar

SM Buffer

1st make: 100 ml of 1 M Tris-HCl (pH 7.5)
into 50 ml mega ohm water add 15.7 g Tris HCl
pH to 7.5
Bring up to 100 ml in volumetric cylinder
2nd make: 2% (w/v) gelatin (meaning 2.0 grams gelatin/100 ml water)
into 80 ml mega ohm water add 2.0 gm EIA purity reagent gelatin
heat to dissolve if necessary
add up to 100 ml in graduated cylinder
3rd make: SM buffer
to 800 ml mega ohm water add –
5.8 g NaCl
2.0 g MgSO4.7H20
50.0 ml 1 M Tris-HCl (pH 7.5)
5.0 ml 2% (w/v) gelatin
add water up to 1 L
filter sterilize into 250 ml or 500 ml jars

10x TBS (Tris-Buffered Saline)

Dissolve:
60.5 g Tris Base (50 mM)
87.6 g NaCl (150 mM)
in 800 ml of D/W
Adjust pH to 7.5 with 1 M HCl (about 9.5 ml)
Add D/W to 1 L
Filter
Store at 4C
TBS is stable at 4C for 3 months

DEPC Treated Water

Use large orange, capped bottles only
Fill to 1400 ml with mega ohm water
Add 1400 ul of DEPC
Shake
Let sit overnight
Autoclave (liquids, 40 min)

X-gal (5-Bromo-4-chloro-3-indolyl-b-D-galactoside)

Dissolve X-gal in dimethyformamide at 20 mg/ml in a glass or polypropylene tube
Wrap tube with aluminum foil
Store at –20°C

0.5M EDTA (pH 8.0)

186.1 g/L in H2O
pH to 8.0 using NaOH pellets.

NOTE: EDTA will not begin to dissolve until it is close to reaching the target pH. Add ~1/3 of the EDTA at a time and wait until it is dissolved (by adding NaOH pellets) before adding more EDTA.

NOTE: It is recommended to make up only 100mL, as this volume is likely to last years.

10/27/2011 SJW


Sterile Water

Rinse used sterile water jar 3 x with mega ohm water
Fill with mega ohm water to about 20 ml in small screw top jar
Autoclave

5 x Formaldehyde Gel Running Buffer (for Northerns)

Dissolve 4.1 g Sodium Acetate into 1 L DEPC water
Take 800 ml and put into new beaker (throw away the remaining 200 ml)
Add 20.6 g MOPS
Adjust to 7.0 with NaOH
Add 10 ml 0.5 M EDTA in DEPC water
Adjust volume to 1 L with DEPC water
Filter
Cover jar with Al foil and store in the dark

8X His Column Binding/Solubilization Buffer (Invitrogen ProBond Resin)


40mM imidazole (2.725 g/L)
4M NaCl (233.6 g/L)
160mM Tris-HCl (25.216 g/L)
16M urea (961 g/L)

10X PBS


NaCl - 80 g/L
KCl - 2 g/L
Na2HPO4 - 14.4 g/L
KH2PO4 - 2.4 g/L

pH = 7.5

1X TBS-T


Dilute 10X TBS (in 4C) to 1X and add 0.05% Tween20 (0.5mL/L)

NOTE: Do not autoclave with Tween20.


Nucleic Acid

Nucleic Acid Precipitation

More Info: "The Basics: How Ethanol Precipitation of DNA and RNA Works @ Bitesize Bio"

Ethanol Precipitation (DNA)

Add 1/10 volume of 3M NaOAC (pH=5.2) to sample.
Invert tube several times to mix.
Add 2 volumes of ice cold 100% EtOH to sample.
Invert tube several times to mix.
Incubate @ -20C for at least 30 mins. Can go longer if desired.
Centrifuge at highspeed (~16,000g) @ 4C for 30 mins to pellet DNA.
Discard supernatant.
Wash pellet with 70% EtOH.
Spin 5 mins. at highspeed (`16,000g) @ 4C to repellet DNA.
CAREFULLY remove supernatant.
Air dry pellet for 5-10 mins.
Resuspend DNA pellet in appropriate volume of water or Qiagen EB (10mM Tris-HCl, pH=7.5)


RNA Extraction [Cost per mL ~$1.10; Cost per sample, including mortar/pestle ~ $2.25]

Manufacturers' Protocol - MRC
  1. Turn centrifuge on to cool to 4C
  2. Clean Homogenizer
- Rinse in DEPC water in 50 ml falcon tube (3x – 3 separate tubes)
  1. Get sample and thaw enough to get out of container
  2. Measure weight of sample
  3. Take sample out (screw out and use forceps) and chop up with sterile razor blade
  4. Put tri-reagent (stays on ice when not using & is light sensitive) into 50 ml falcon tube (or smaller tube depending on size of sample) – for a 0.7 gram sample I used 7 ml of tri-reagent. Note: in 50 ml falcon tube need at least 3 ml of tri-reagent to get it to work
  5. Add sample
  6. Keep on ice
  7. Blot homogenizer with paper towel to remove excess water
  8. Homogenize sample (don’t leave off ice for too long)
  9. Homogenize until sample is in solution
  10. Transfer all or part (I kept 6 ml) of mixture into a 13 ml tube (only add up to 7 ml)
  11. Let sit for 5 min at RT
  12. Rinse homogenizer in DEPC water (same tube used to clean in the beginning)
  13. Add 0.2 ml of chloroform (under hood, open only in hood, pour into glass beaker first) per 1 ml of tri-reagent
  14. Cover & shake
  15. Let sit for 15 min at room temp
  16. Change gloves
  17. Spin at 12,000 x g (11,500 rpm) for 15 min at 4C
  18. Transfer aqueous (top) phase to fresh tube (top layer has the RNA in it – bottom layer has DNA and proteins in it)
  19. Add 3 ml iso. (2 – Propanol, under hood) to precipitate out the RNA
  20. Cap and vortex
  21. Let sit at RT for 10 min
  22. Put waist with tri-reagent etc. in tri-reagent bottle in fume hood
  23. Clean up homogenizer and put away (put in 50 ml falcon tube with 5-7 ml 30% H2O2 and up to 40 ml with DEPC water
  24. Spin at 12,000 x g (11,500 rpm) for 15 min at 4C
  25. Remove supernatant (I want the pellet – RNA)
    1. Get glass beaker and paper towels (small stack)
    2. Pour off supernatant into beaker and place tube upside-down on paper towels
    3. Note: do not rock the tube back and fourth or will loosen pellet
  26. Add 1 ml 75% EtOH in DEPC water per 1 ml of tri-reagent added in beginning
  27. Cap & move – rock back and fourth to loosen pellet – vortex if necessary
  28. Spin 11,500 x rpm for 5 min at 4C
  29. Remove supernatant again & put on paper towel – be much more careful to make sure pellet does not slip out
  30. Spot Spin – turn on centrifuge, let go up to about 1000 rpm then shut off – note: place pellet facing upward
  31. Use filter pipette tips to remove excess EtOH
  32. Turn upside down on paper towels
  33. Wait 10 mins
  34. Depending on size of pellet add dnase free water to the tube – if taking to mRNA always use 500 ul (I used 500 ul for the ovary – large pellet, and 250 ul for the muscle)
  35. Dissolved into solution by pipetting
  36. Put in 1.5 ml tube
  37. Put on ice
  38. Spec to determine how much RNA you have

Updated cost 11/9/2012 SJW


mRNA Isolation (Progmega IV)


  1. Place 500 ul of sample or diluted sample (you do not want more than 1000 ug) in 65C heat block for 10 mins
  2. Add 3 ul Oligo dT probe & 13 ul of 20 x SSC to the RNA (be careful not to let tubes cool at all)
  3. Vortex
  4. Incubate at RT until completely cooled (about 10 min)
  5. In the meantime make stocks
    1. Prepare 1.2 ml of sterile 0.5 x SSC (for 1 rxn use):
i. 30 ul of 20 x SSC
ii. 1.170 ml of RNase free water
    1. Prepare 1.4 ml of sterile 0.1 x SSC (for 1 rxn use):
i. 7 ul of 20 x SSC
ii. 1.393 ml of RNase free water
  1. Resuspend one tube of SA-PMP per isolation by gently flicking the bottom of the tube until completely dispersed
  2. Put tube in magnetic stand
  3. Remove supernatant with thin glass pipet – careful not to touch particles
  4. Wash the SA-PMPs 3 x with 0.5 x SSC – use 300 ul per wash and completely resuspend particles during each wash
  5. After the last wash try to remove all liquid possible
  6. Resuspend the washed SA-PMPs in 100 ul of 0.5 x SSC
  7. Add entire sample mixture to the tubes containing the washed SA-PMPs
  8. Incubate at RT for 10 min; gently mixing by inverting every 1-2 min
  9. Put in magnetic stand
  10. Wash the particles 4 x with 0.1 x SSC
  11. Resuspend in 100 ul of RNase free water and gently resuspend by flicking tube
  12. Put in magnetic stand & transfer the eluted mRNA to a sterile RNase free tube (1.5 ml). Do not throw particles away
  13. Repeat elution step by resuspending in 150 ul of RNase free water; repeat capture step, pooling the elute with the mRNA eluted in the last step
Precipitate out mRNA
  1. Add a tenth volume of 2M Sodium Chloride in DEPC treated water (25 ul)
  2. Mix
  3. Add 4 volumes (1 ml) of 100% EtOH
  4. Vortex
  5. Put in –20C overnight
  6. Spin @ 4C for 15 min at top speed; mark tube so that can find pellet
  7. Decant supernatant; twist cap behind tube and put upside-down on paper towels then blot
  8. Wash with 75% EtOH in DEPC treated water (use about 1 ml); rock back and fourth very carefully 2-3 times
  9. Spin at 4C for 5 min at top speed
  10. Decant and blot (same way as above)
  11. Repeat wash
    1. Add 1 ml 75% EtOH in DEPC treated water
    2. Spin 4C for 5 min at top speed (go longer if pellet loosens)
    3. Decant and blot
  12. Spot spin to collect remaining liquid
  13. Remove all fluid (use filtered med or sm filtered tip to drag fluid up to dry)
  14. Dry for a variable amt of time (usually 10 min but don’t want to dry out)
  15. Resuspend
    1. If going to do a northern add 10 ul formazol to pellet (store at –20C)
    2. If going to do RT PCR or a library then add 10 ul RNase free water (store at –80C)
  16. Flick pellet until totally dispersed
  17. Freeze until ready to use
  18. Spin at top speed for 5 min at 4C
  19. What you want is the supernatant
  20. Keep on ice
  21. Spec to determine how much RNA you have use:
    1. 0.4 ul sample
    2. 99.6 ul water

Reverse Transcription (Promega M-MLV: Cat#M1701; ) [Cost per sample ~$1.50]


A single reaction volume = 25uL. The volume of RNA, primer(s) and M-MLV RT used are variable and will be specific to your current experiment. The directions below apply to a reaction using 1ug of total RNA. You may need to make changes to accommodate your own conditions.

  1. Use as much RNA as possible (up to 1ug); max volume of RNA = 17.75uL. Generally, identify the RNA sample with the lowest concentration and multiply by 17.75uL. Use this quantity (ug) of RNA for each and every sample.
  2. Transfer calculated volume(s) of RNA to 0.5mL snap cap tubes or PCR plate. Adjust volumes of individual samples to 17.75uL with H2O.
  3. Add appropriate amount of primer to sample. Use 0.25ug primer per 1ug of RNA in sample (= 0.5uL of Promega oligo dT Cat#C1101 in this example). Total volume (RNA + primers) should equal 18.25uL.
  4. Heat samples at 70C for 5 min in thermocycler.
  5. Place samples on ice IMMEDIATELY.
  6. Make Master Mix:

PER RXN
5 uL 5x Buffer (M-MLV RT Buffer)
1.25 uL 10mM dNTPs (Promega Cat#U1511)
0.5 uL M-MLV RT per ug of RNA

7. Mix well.
8. Add 6.75uL of master mix to each reaction.
9. Mix well, but do not vortex.
10.Spot spin.
11.Incubate @ 42C for 1hr in thermalcycler for oligo dT primers OR @ 37C for random primers.
12.Heat inactivate @ 95C for 3 min.
13.Spot spin.
14.Store @ -20C.

3/17/2011 SJW

Quantitative PCR (qPCR)/Real-time PCR (2x Sso Fast EvaGreen Supermix) [Cost per rxn ~$0.42]

Single reaction (20uL) set up is listed below. Be sure to make a master mix volume that will accommodate the following: all of your samples, two water (no template controls; NTC) samples, plus an extra 10% to accommodate pipetting errors. Distribute appropriate amount of master mix (volume of master mix + template = 20uL) to white PCR plate. Add template. Cap with optical PCR caps. Spin plate for 1min @ 3000g. Put in thermalcycler.

Reaction_Components
Volume
Final Concentration
2x Sso Fast EvaGreen
10
1x
Forward Primer (10uM)
0.5
0.2uM
Reverse Primer (10uM)
0.5
0.2uM
Template
Up to 5uL

H2O (PCR grade)
variable
Use to bring reaction volume up to 20uL

See the BioRad Sso Fast EvaGreen Supermix information sheet for cycling parameters, as they are dependent upon template (cDNA or gDNA).


3/17/2011 SJW


Conventional PCR (2x Apex Red) [Cost per rxn ~ $0.52]

Single reaction (25uL) set up is listed below. Be sure to make a master mix volume that will accommodate all of your samples, two water (no template controls; NTC) samples, plus an extra 10% to accommodate pipetting errors. Distribute appropriate amount of master mix (volume of master mix + template = 25uL) to PCR tubes or PCR plate. Make sure all tubes/caps are tightly closed. Put in thermalcycler.
Reaction_Components
Volume
Final Concentration
2x Apex Red
12.5
1x
Forward Primer (10uM)
0.5
0.2uM
Reverse Primer (10uM)
0.5
0.2uM
Template
Up to 5uL

H2O (PCR grade)
variable
Use to bring reaction volume up to 25uL
Typical cycling paramaters (ask for help on using the thermal cycler):

95C - 10mins
40 cyles of:
95C - 15s
50-60C - 15s
72C - 10s - 2mins (dependent on amplicon size; ~1000kb/min)

3/17/2011 SJW

Conventional PCR (2x GoTaq Green)

Polymerase Chain Reaction involves amplifying a DNA (genomic or complementary) target using a polymerase, primers (short oligonucleotide), and dNTPs (A, C, T, Gs). In general the reaction is placed in a machine (thermocycler) where a series of temperature changes are performed [Denature (~94C), Anneal (primer specific ~50-60C), and Extention (~72C)].
For this lab you will be using Promega's GoTaq Product. Please Read!

Prepare your samples in duplicate AND make sure to include at least 2 negative controls for each primer (no template).

For a 50μl reaction volume:
Component
Volume
Final Conc.
GoTaq®Green Master Mix, 2X
25
1x
upstream primer, 10μM
0.5–5.0μl
0.1–1.0μM
downstream primer, 10μM
0.5–5.0μl
0.1–1.0μM
DNA template
1–5μl
<250ng



Cloning (pCR 2.1 TA)

  1. Heat water bath to 42C
  2. Determine what pieces you want to clone
  3. Let band thaw @ RT
  4. Put in Ultrafree-DA (in my area right of empty boxes)
  5. Spin at 5000 g for 10 min at RT
  6. Thaw SOC Medium which is at –80C 3rd draw down in bucket
  7. Use variable amount – 0.5 to 4 ul for next step, if use 4 ul then do:
    1. 4 ul PCR product (take out)
    2. Add 1 ul Salt Sol
    3. Add 1 ul Vector
    4. Total 6 ul
  8. Incubate for 10-15 min at 22C in thermocycler (no vortexing, etc.)
  9. Directly to ice
  10. Take 2 ul Cloning RXN mixture and add to competent cells (purple cap), do not vortex; do not pipet up and down
  11. Incubate on ice 10 min
  12. Heat shock for exactly 30 sec at 42C – NO shaking (use water bath)
  13. Put on ice for 2 min
  14. Add 250 ul ROOM TEMP SOC Medium
  15. Cap (purple lid) and roll so that mixture coats tube
  16. Put in 37C incubator @ 225 rpm for 1 hr – laying sideways
  17. 30 min into incubation take out Kanamycin Plate to get to RT (can put in 37C incubator – no shake)
  18. When plate is up to room temp, spread 40 ul x-gal on plate then dry plate at 37C
  19. Plate out some of the mixture (with competent cell in it), can use 100 ul and save the rest at 4C
  20. Leave upside-down in incubator at 37C overnight
  21. Pick out colonies the next day
  22. Streak out the colonies that you want onto a fresh plate – make grid on plate– don’t over do, just touch colony with yellow tip and streak into a Z shape (should streak out at least one blue colony as well)
  23. Let plates grow up overnight – or if do 1st thing in the morning may have colonies by late afternoon
  24. Do Colony PCR (per rnx)
    1. RNase free water 43.4 ul
    2. Buffer A (fisher) 5 ul
    3. dNTPs 1 ul
    4. Primer F (M13) 0.2 ul
    5. Primer R (M13) 0.2 ul
    6. Taq 0.2 ul
    7. Touch loop to plate and then swirl in MM
    8. Run in thermocycler under SBR, CLNY
    9. Run one positive control and a blue colony
    10. Run on gel
  25. Inoculate Cultures
    1. Touch plate and put into and swirl around in a 50 ml falcon tube with about 30 ml of LB broth in it
    2. Incubate at 37C (shake) overnight
  26. Mini Prep
    1. Next morning, spin down for 10 min @ 4C @ 3500 rpm
    2. Decant
    3. Turn upside down on paper towel for 5 min, blot a few times
    4. Put in –20C freezer or add 250 ul cell resuspension solution, vortex, then put in –20C freezer
    5. Move to screw cap tube or regular centrifuge tube (1.5 ml)
    6. Add 250 ul Cell Lysis Solution to each sample
    7. Invert 4-6 times – be gentle
    8. Wait 5 min at RT
    9. Set up timer to 5 min for next step
    10. Add 10 ul of Alkaline Protease Solution (do very quickly so that all samples are done at about the same time)
    11. Invert 4 times to mix
    12. Let go for 4 min but NO longer than 5 at RT
    13. Add 350 ul Neutralization Solution
    14. Invert 4-6 times
    15. Spin at top speed 10 min at RT
    16. Set up Column
    17. Decant Cleared lysate into spin column
    18. Spin at top speed for 1 min at RT, Discard flow through
    19. Add 750 ul wash solution (with EtOH)
    20. Spin top speed for 1 min, Discard flow through
    21. Add 250 ul wash solution (with EtOH)
    22. Spin 1 min, Discard flow through
    23. Spin 2 min RT
    24. Transfer spin column to 1.5 ml tube (cut off cap)
    25. Add 100 ul Nuclease-free water
    26. Spin top speed 1 min at RT
    27. Spec or Store at –20C
    28. Spec in microcell – use 5 ul sample + 95 ul water (wavelength 260)
    29. Sequence


Northern Blotting


Night before running gel
  1. Clean out gel box
    1. In DEPC treated water jar mix 75 ml 30% H2O2 up to 1 L with DEPC treated water
    2. Put in medium gel box with everything will need next day (combs & tray)
  2. Make sure glassware & DEPC water is ready
    1. Cylinder, orange capped bottles, filter tips, water bath, gloves, etc.
  3. Prepare samples
    1. Spec
    2. Determine how much sample to use to get 1 ug of mRNA
Day 1: Running Gel

  1. Turn on water bath to 60C
  2. Turn heat block to 55C
  3. Prepare agarose
    1. 1.2 g agarose (over with other chemicals by Steven – use for RNA only)
    2. 75 ml DEPC water
    3. lightly cap
    4. heat in microwave about 2-3 min swirling frequently
    5. let cool a little bit
    6. put in 60C waterbath
  4. Rinse gel box
    1. Use one jar of DEPC treated water to wash down everything
    2. Put gel box, tray, and combs on Al foil until ready to use
  5. Move equipment to hood
    1. Large gel box – level it
    2. Med gel box on top of large gel box
    3. Power supply
  6. Add solutions to gel
    1. Add 22 ml formaldehyde (near gel box area in cabinet) – use 25 ml pipet and swirl once added
    2. Add 24 ml 5 x Running Buffer – use 25 ml pipet and swirl once added
  7. Pour gel
    1. Look first to make sure homogenous and no dirt in the mixture
    2. Get out any air bubbles right away with filter tip
    3. Close hood
    4. Turn water bath down to 55C
    5. Clean bottle that had gel with soap and water
  8. Set up samples
    1. Let samples thaw, if mRNA homogenize completely, vortex, and spin for 5 min
    2. Prepare loading buffer
i. 159 ul RNA solution (Sample Buffer)
ii. 81 ul formaldehyde
iii. mix in RNase free 1.5 ml tube
iv. vortex
v. spot spin
    1. Prepare samples & marker
i. Add appropriate amt of formazol (want total with sample will be 10 ul)
ii. Add appropriate amt (1 ug) of sample
iii. Add 10 ul loading buffer
iv. Don’t forget to prepare positive control
1. can use 56 mRNA cut ovary (.84 ul sample + 9.16 ul formazol)
v. vortex
vi. spot spin
vii. for marker want 5 ul marker, 5 ul formazol, 10 ul loading buffer, 1 ul EtBr)
viii. Denature for 15 min (at least) in 55C heat block (all samples including marker)
  1. Make 1 x Running Buffer
    1. Make in empty DEPC water and label, use sterile graduated cylinder)
    2. 160 ml 5 x Running Buffer (found in same place as formaldehyde)
    3. 640 ml DEPC treated water
    4. Shake
  2. Prepare to load gel
    1. Take out comb
    2. Turn gel the right way
    3. Add all of the running buffer
    4. Put cover on
    5. Turn on voltage to 70 for at least 5 min or until ready to load
  3. Add 1 ug of EtBr to total RNA sample BUT NOT to mRNA samples
    1. Vortex
  4. Spot spin all samples
  5. Load 20 ul sample – be sure to try to add exactly equal amounts in all wells
  6. Write down the order of the samples
  7. Run at 120 volts until dye reaches 7 cm (about 2 hr and 10 min) then run at 55 volts for 1 cm (about 30 min) (takes about 3 hr total)
  8. Expose to UV if have total RNA samples with EtBr & take picture (don’t expose to UV light too long)
  9. Put in plastic tray
  10. Put DEPC water to cover – swish and pour out
  11. Cover again with DEPC water; shake 30 min changing water 3-4 times
  12. Get 14 cm x 11 cm membrane (under saran wrap and Al foil in Dimi’s area)
  13. Stack 20 sheets of GB004; then 4 sheets of GB002
  14. Get and additional 1 GB002 and wet in 20 x SSC and put on top of stack
  15. Get air bubbles out
  16. Soak membrane in DEPC water (just swish around), then in 20 x SSC for 5 min
  17. Put membrane on stack
  18. Get rid of any air bubbles
  19. Cut last lane off of get (won’t fit unless you cut one lane off)
  20. Put gel on paper – must put it on right the first time, can not pick back up and put on again
  21. Wet 3 GB002 in 20 x SSC and put over gel; make sure to get air bubbles out
  22. Put buffer tray on
  23. Put wick paper on and fold over to reach into tray
  24. Put wick cover on
  25. Fill upper tray with 20 x SSC
  26. Cover carefully with Al foil
  27. Leave overnight

Day 2: Washing and Hybridization

  1. Wash membrane twice in 100 ml 2 x SSC (20 ml 20 x SSC up to 200 ml with DEPC water)
    1. Use clear tray over by gel box
    2. Uncover membrane
    3. Throw everything up to gel away (except for cover)
    4. Mark where lanes of gel are on membrane
    5. Take gel and look to make sure there are bands (if used EtBr) if did not use EtBr then don’t do it
    6. Look at membrane under UV box and mark with pencil where the bands are (use clean forceps when moving membrane)
    7. Move to other UV box and take a picture of membrane with a ruler that lines up along the side starting at the wells
    8. Put membrane into tray
    9. Wash twice with 100 ml of 2 x SSC for 5 min each
  2. Cut 2 pieces of filter paper enough to cover membrane
  3. Put membrane on one piece of filter paper (above 20C freezer by Steven’s area)
  4. Crosslink it (crosslinker, on, 1200, start; connects RNA to membrane)
  5. Put second filter paper over membrane
  6. Wrap with saran wrap and label
  7. Put at 4C until needed
  8. To make probe and run pre-labeling rxns it’s a good idea to use a plasmid prep as your template and not cDNA
  9. Run pre-labeling rxn to determine the conditions to run labeling mix at – do everything the same except use regular dNTPs. Optimize this PCR rxn before making probe. May need to change the following protocol (MM and running conditions) as necessary.
  10. Using degenerative primers to make probe is okay as long as you get a clean band
  11. In the meantime, make probe (1 reaction) Change primer and template as necessary
RNase free water 38.6 ul
Ampli Buffer 5.0 ul
DNTPs (Labeling Mix –20 blue/black box) 8.75 ul
Primer cSTAR F1 0.4 ul
Primer cSTAR R1 0.4 ul
Ampli Taq Polymerase 0.4 ul
Template (cSTAR 8 Clone) 0.2 ul
Magnesium Chloride (25 mM) 2.5 ul
*this primer pair will isolate about 500 base pairs
Vortex
Spot spin
Put in DNA Engine – Labeling
1 = 94 for 5 min
2 = 94 for 1 min
3 = 50 for 1 min
4 = 72 for 2 min
cycle 35 x, run, yes heat lid
can reuse probe because only use 3 ul for each northern (store at 4C)
  1. Run 5 ul probe on gel and take picture to make sure that it worked and have a strong band
  2. Dimi does not quantify the probe – just make sure you have a strong band before continuing
  3. When probe is ready can continue on (need to run on gel and see if worked – 685 bp)
  4. Put hybridizing machine (near crosslinker) at 42C
  5. Get Calf Thymus DNA (sigma) from –20C (used to prevent non-specific binding)
    1. Thaw out
    2. Take 150 ul and put in screw cap 1.5 ml tube (will use only 110 ul if using 10 ml hybridization buffer)
    3. Boil in glass beaker over flame for 5 mins
    4. Move directly to ice when done & use when cool
  6. Get Formamide Hybridization Buffer from 4C fridge and put 10 ml into a 50 ml falcon tube; put in hybridization machine to warm – should became clear
  7. Get membrane and cut off dye front
  8. Get red capped jar and put membrane in (the long way, don’t overlap, put RNA side to the inside)
  9. Cap and put in hybridization machine at 42C
  10. Add 110 ul Calf Thymus DNA to hybridization buffer
  11. Mix well
  12. Pour in to jar with membrane
  13. Roll so that solution gets all the way around
  14. Don’t want any air bubbles
  15. Put in hybridization machine with rotate on
  16. Leave alone for 1 hr
  17. 10 min before ready to put probe in (50 min later) denature probe in thermocycler for 10 min at 95C
  18. Put directly on ice after 10 min
  19. Take a few ml of liquid in jar and put in 50 ml tube, then add 3 ul probe directly to it, mix very well and then put back in jar, and put jar back in hybridization machine (use 3-5 ul probe per 10 ml of hybridization buffer)
  20. Leave alone for 16-18 hr
Day 3: Washing and Viewing Northern

  1. Washing may vary depending on the size of the probe you use. This protocol is using high stringency (low salt and high temp) and the probe is about 500 bp long. If the probe is shorter use a less stringent wash (higher salt with lower temp)
  2. Warm Shake and Bake oven for at least 20 min at 70C (can vary temperature – the higher the temp the lower the background but it gets rid of some of the signal, 70 is a high temp, the protocol calls for 55C)
  3. Warm heat bath to 55C
  4. Make two washing solutions
    1. 0.2 x SSPE
i. need 100 ml per wash and need 4 washes
ii. put 4 ml 20 x SSPE in jar labeled 0.2 x SSPE
iii. add 376 ml DEPC water to it
iv. shake
v. add 20 ml 10% SDS (stored near 20 x SSPE)
vi. pre-warm wash for a few min in microwave
    1. 1 x SSPE
i. 10 ml 20 x SSPE
ii. 190 ml DEPC water
iii. Put in hybridization incubator at 70C
  1. Get clear tray
  2. Take jar out of hybridization incubator
  3. Take membrane out carefully with forceps and put RNA side up on the tray (can save what’s in the jar)
  4. Add a small amount of 0.2 x SSPE, swish, and drain
  5. Add 100 ml of 0.2 x SSPE
  6. Put in Shake and Bake oven on max speed for 15 min
  7. Put 0.2 x SSPE in hybridization incubator at 70C
  8. Drain, repeat with fresh wash (100 ml, 15 min)
  9. Drain, repeat wash but 100 ml for 30 min
  10. Drain, repeat wash again 100 ml for 30 min
  11. Drain, wash with 100 ml 1 x SSPE for 5 min
  12. Drain, repeat wash with 1 x SSPE
  13. In the meantime make solutions
    1. Blocking Solution (blocking bottle)
i. 20 ml Detector Block Solution (5 x; at 4C)
ii. add 80 ml DEPC water
iii. shake slowly by inverting
iv. add 0.3 g (300 mg) Detector Blocking Powder
v. shake
vi. warm in water bath at 55C for about 20 min or until powder is in solution
vii. once in solution leave at RT (DON’T use until at RT, put in 4C fridge if necessary)
    1. Phosphatase Wash (wash bottle)
i. 60 ml phosphatase wash solution (5 x; at RT; need to warm at 37C before using to get everything into solution)
ii. 240 ml DEPC water
iii. shake
    1. Assay Solution (assay bottle)
i. 10 ml phoshatase assay buffer (10 x; at 4C)
ii. 90 ml DEPC water
  1. After washes are complete put membrane (get drippings off first) into pink case and add 50 ml blocking solution (use exactly 50)
  2. Incubate for 45 min at RT (leave door open) in Shake and Bake oven (with shake on)
  3. After 45 min add 5 ul of enzyme (alcholine phosphatase) to blocking bottle with 50 ml blocking solution in it
  4. Mix well
  5. Remove blocking solution from pink tray
  6. Add 50 ml blocking solution plus enzyme
  7. Incubate at RT for 30 min in Shake and Bake oven
  8. Transfer membrane to bigger tray
  9. Wash with 100 ml phosphatase wash for 5 min in Shake and Bake oven
  10. Drain
  11. Repeat wash for 5 min two more times (total of 3 washes)
  12. Remove wash
  13. Transfer membrane to clean clear tray
  14. Add 50 ml assay to membrane to neutralize
  15. Swish around
  16. Leave on 10 min, changing assay once after a few min
  17. Get metal lid from transfer system
  18. Moisten and cover with saran wrap
  19. Take out membrane and blot on filter paper (both sides)
  20. Lay membrane on transfer system lid (RNA up) and get rid of any air bubbles
  21. Cover with 7-8 ml of CDP-Star4 (100 ml ready to use)
  22. Leave on 5 min
  23. Remove liquid and blot onto filter paper (just like with western)
  24. Put membrane on and cover with saran wrap (like western)
  25. Expose same way you do western – can try 1 min, 10 min, then 1 hr


Protein

Extraction

  1. Thaw samples on ice
  2. Add 300 ul of Cell Lytic
  3. Smash up with pipet tip
  4. Vortex
  5. Spin 15 min @ max speed @ 4C
  6. Remove supernatant into fresh tube (but keep debris at –20C)
  7. Take 50 ul of supernatant and put in fresh tube
  8. To the 50 ul add 1.5 ml of Coomassie Plus – 200 Protein Assay
  9. Mix
  10. Spec in disposable 2.0 ml cell @ wavelength 595
  11. Want a reading between 0.4 and 2.0
  12. Dilute sample and rerun if necessary
  13. Determine how much protein you have and calculate how much water you need to add to 50 ul to get your appropriate concentration (1 ug/ul) (use protein specs spreadsheet – in my documents in specs folder)

Run Proteins on Gel (Pierce 4-20% Tris-Hepes)


1. Start boiling water.
2. Aliquot appropriate volume (no more than 25uL) of sample in to SCREW CAP tube.
3. Combine sample with equal volume of 2x sample reducing buffer.
4. Boil 5 minutes. Conduct steps 5-10 while boiling (not a requirement; just convenient).
5. Get gel(s) (in 4C).
6. Open package and install into gel box.
7. Get 500mL of 1x tris-hepes running buffer.
8. Fill inner chamber with 1x tris-hepes running buffer. Ensure that there are no leaks.
9. Pour remainder of buffer into outer chamber.
10. Rinse each well 3 times with clean gel loading tip.
11. Spot spin samples.
12. Load 10uL of Invitrogen SeeBlue protein standard (Current as of: 20081030)
13. Load samples into wells.
14. Run gel @ 150V ~40mins.
15a. Proceed to Western Blotting
OR
15b. Stain gel in Coomassie stain for 45 mins. with shaking.
16. Pour Coomassie stain back into bottle.
17. Rinse gel briefly with 10% acetic acid and discard liquid.
18. Destain gel with 10% acetic acid with shaking until bands are visible. Requires periodic changes of destaining solution. Usually takes 2-3hrs to start seeing bands.

Western (generic nitrocellulose)

NOTE: This is a DOWNWARD trasfer system. Currently using Millipore Immobilin chemiluminescent developers; need 3mL of each reagent (20081030).
1. Equilibrate gel and nitrocellulose (separate containers) in transfer buffer 5mins.
2. Get thick filter paper (need 2 per gel) and cut to correct size.
3. Quickly dip into transfer buffer with thin filter paper in it (don’t wet too much).
4. Wet down transfer system with wet paper towel.
5. Place thick filter paper on transfer system (remove air bubbles).
6. Put notch on thin membrane and place on thick filter paper (remove air bubbles).
7. Place gel over membrane (remove air bubbles).
8. Put large filter paper over gel.
9. Remove air bubbles.
10. Get rid of excess fluid with Kim wipe.
11. Put metal then plastic cover on.
12. Run at 15 volts for about 15 min. May want to run longer if target is large molecular weight (>50kDa).
13. In the meantime make blocking solution – need 60 ml of blocking solution per gel Use 1.5 g dry milk per 30 ml 1x TBS-T. Make in beaker and put on stir plate.
14. Stain gel if necessary see above.
15. Put filter and 30 ml of blocking solution into small container.
16. Shake slowly for at least 1 hr (could go overnight if need to).
17. Add appropriate volume of primary antibody. Will vary depending on antibody to be used. See antibody list for appropriate dilution.
18. Shake slowly for at least 1 hr (could go overnight if need to).
19. Wash - Drain milk mixture.
20. Add small amount of 1x TBS-T. Shake a bit and pour out.
21. Add 1 x TBS-T to cover filter paper well.
22. Shake 3 x 10 mins.
23. Add 30 ml of blocking solution.
24. Shake for 5 min.
25. Add appropriate volume of secondary antibody conjugated to horseradish peroxidase. See antibody list for appropriate dilution.
26. Shake for about 30 min.
27. Pour out blocking solution and rinse out with 1x TBS-T.
29. Remove developing solutions from 4C to allow to equilibrate to room temp.
30. Add enough 1 x TBS-T to wash. Wash 3 x 10mins.
31. If taking membrane to another lab for imaging, gather the following items: Membrane in 1x TBS-T, plastic wrap, 2 x 15mL conicals containing appropriate volumes of unmixed developing solutions, serological pipette (10mL), gloves, paper towel or two.
32. Take off wash and get semi-dry (blot on filter paper).
33. Place saran wrap down on flat surface.
34. Combine two developing solutions, mix and pour/pipette onto membrane.
35. Wait 5 min.
36. Remove membrane and blot on paper.
37. Place membrane face down on a NEW piece of plastic wrap.
38. Image blot. ASK FOR DIRECTIONS ON HOW TO IMAGE BLOT.
39. Exposure times will vary. Currently, 5-10mins seems to be good for most antibodies (20081030).

Run Proteins on Gel (NuPAGE)

More Info: "How SDS-PAGE works @ Bitesize Bio"
  1. Turn on heating block to 70C or set thermocycler to 70C
  2. Aliquot out 13 ul of diluted protein sample into fresh tube
  3. Add 4 x (5 ul) loading die buffer (NuPAGE LDS Sample Buffer / blue-purple color)
  4. Add 10x (2 ul) reducing agent (NuPAGE Sample Reducing Agent / green cap)
  5. Get appropriate gel(s) – careful liquid is nasty
  6. Rinse off gel(s) with distilled water
  7. Peal off white sticker
  8. Take out comb
  9. Face lower part of gels to each other
  10. Fill in between gels with 1 x MES Buffer
  11. Get rid of bubbles
  12. Clean out each well 3 times with long narrow tips
  13. Denature proteins for 10 min at 70C in heat block or thermocycler
  14. Add 500 ul NuPAGE Antioxident to inner chamber
  15. Note: if not running a reducing gel then do not add Reducing agent or antioxident
  16. Fill rest of chamber with 1 x MES (about ¾ full)
  17. Fill up rest of inner chamber with 1 x MES
  18. After samples have been heated for 10 min spot spin them at room temp
  19. Load 3 ul of marker (orange cap @ -20C)
  20. Load 3 ul of each sample (can reuse tip if rinse between each sample)
  21. Run at 200 volts for about 30-40 min
  22. Store remaining sample @ -20C
  23. After run is complete, crack open gel
  24. Remove lanes
  25. Remove ridge
  26. Carefully remove gel
  27. Go onto western or directly stain
  28. To stain: pour small amount of Coomassie Stain in small container, add gel, and shake slowly
  29. Leave for a while (hour or more)
  30. To destain, pour stain back into original container, put small amount of 10% acetic acid over gel, swirl around, dump out, add more 10% acetic acid, let shake slowly for a while (until can see bands)

Western (NuPage)

  1. Soak thin paper in 2x NuPAGE Trans with 10% Methanol and 1:1000 NuPAGE Antioxident for about 15 min slowly on shaker
  2. Get thick filter paper (need 2 per gel)
  3. Quickly dip into buffer with thin filter paper in it (don’t wet too much)
  4. Wet down transfer system with wet paper towel
  5. Place thick filter paper on transfer system (remove air bubbles)
  6. Put notch on thin membrane and place on thick filter paper (remove air bubbles)
  7. Place gel over membrane (remove air bubbles)
  8. Put large filter paper over gel
  9. Remove air bubbles
  10. Get rid of excess fluid with Kim wipe
  11. Put metal then plastic cover on
  12. Run at 15 volts for about 15 min
  13. In the meantime make blocking solution – need 60 ml of blocking solution per gel
Use 1.5 g dry milk per 30 ml 1 x TBS-T
Make in beaker and put on stir plate
  1. Stain gel if necessary see above
  2. Put filter and 30 ml of blocking solution into small container
  3. Shake slowly for at least 1 hr (could go overnight if need to)
  4. Add 30 ul of primary antibody
  5. Shake slowly for at least 1 hr (could go overnight if need to)
  6. Wash - Drain milk mixture
  7. Add small amount of 1 x TBS-T
  8. Shake a bit and pour out
  9. Add 1 x TBS-T to cover filter paper well
  10. Shake (about 3) for about 30 min
  11. Change wash again in 15 min, then every 5 min for 15 min
  12. Remove wash
  13. Add 30 ml of blocking solution
  14. Shake for 5 min
  15. Add 2 ul of Anti-Rabbit Ig Horseradish Peroxidase
  16. Shake for about 30 min
  17. Pour out blocking solution and rinse out with 1 x TBS-T
  18. Add enough 1 x TBS-T to wash (washing time can vary – ex: wash every 5 min for 20 min, or wash after 15 min then every 5 for a total of 30 min)
  19. Take off wash and get semi-dry (blot on filter paper)
  20. Moisten metal surface on transfer system
  21. Place saran wrap down and try to get rid of the bubbles
  22. Pour 4 ml of mixture over filter
    1. 2 ml Chemi Glow West Luminol/Enhancer Sol.
    2. 2 ml Chemi Glow West Stable Peroxide Buffer
  23. Wait 5 min
  24. Take out filter
  25. Blot on paper
  26. Put between two pieces of plastic or saran wrap and get air bubbles out
  27. To look at it:
    1. Put in gel box on top shelf
    2. Aperture all the way open
    3. Filter 1 on – chemiluminescence
    4. Normal/High
    5. Display Chem
    6. Noise Reduction
    7. 1 min, chemi display, acquire image, expose longer if necessary


RNA to Sequencing Flow Chart

Organism
Extract Tissue
Extract Total RNA
Extract mRNA
Reverse Transcribe to cDNA
Run on Gel
Cut Out Bands
Clone (Clone, Colony PCR, Inoculate, Mini Prep)
Determine Which Colonies to Sequence
Sequence

Mitochondrial Removal
Tissue samples will be homogenized in lysis buffer and mitochondria will be separated from this mixture by differential centrifugation using the Mitochondria Isolation Kit (Pierce, IL, cat no 89874) following the manufactures instructions. Modifications to this protocol will be made in order to isolate the nuclear fraction by continuing the isolation procedure with the pellet formed by centrifugation at 700 x g. Total RNA will then be extracted from these fractions (separately) using Tri Reagent (Molecular Research Center, Inc, Ohio). cDNA libraries will then be constructed from total RNA using the Creator SMART cDNA Library Construction Kit (Clontech, California, cat no K1053-1).

SMART cDNA Synthesis (MassEx)

Note: For “Mass Excision”
Day 1
First Strand cDNA Synthesis
Chill all components and tubes on ice before starting
  1. Combine
1-3 ul RNA Sample (0.025-0.5 ug mRNA or 0.05-1.0 ug total RNA)
1 ul SMART IV Oligonucleotide
1 ul CDS III/3’ PCR Primer
X ul Water up to 5 ul
  1. Mix contents and spot spin
  2. Incubate at 72C for 2 min
  3. Cool on ice 2 min
  4. Spot spin
  5. Add the following
2 ul 5x First-Strand Buffer
1 ul DTT (20 mM)
1 ul dNTP Mix (10 mM)
1 ul PowerScript Reverse Transcriptase
10 ul Total Volume
  1. Mix by gently pipetting
  2. Spot spin
  3. Incubate 42C, 1 hr in an air incubator (says to cover with mineral oil if using thermocycler or waterbath for the incubation)
  4. Put tube on ice to terminate 1st strand rxn
  5. Can stop at this point and leave at –20C if necessary
  6. If continuing on, take 2 ul of 1st strand rxn and put into prechilled 0.5 ml tube
  7. Store remaining 1st strand rxn at –20C for up to 3 months
cDNA Amplification by LD PCR
  1. Preheat thermocycler to 95C
  2. Look at table below to determine how many cycles to run
#
Total RNA (ug) Poly A+ RNA (ug) Number of Cycles
1.0-2.0 0.5-1.0 18-20
0.5-1.0 0.25-0.5 20-22
0.25-0.5 0.125-0.25 22-24
0.05-0.25 0.025-0.125 24-26

  1. Combine
2 ul First-Strand cDNA
80 ul Water
10 ul 10X Advantage 2 PCR Buffer
2 ul 50X dNTP Mix
2 ul 5’ PCR Primer
2 ul CDS III/3’ PCR Primer
2 ul 50X Advantage 2 Polymerase Mix
100 ul Total Volume
  1. Mix contents by gently flicking tube
  2. Spot spin
  3. Start thermocycling using the following cycle
95C 20 sec
cycle X times (based on table)
95C 5 sec
68C 6 min
  1. In the meantime prepare a 1.1% agarose/EtBr gel
  2. Run 5 ul PCR product with a 1 kb marker – the ds cDNA should appear as a 0.1-4 kb smear with a few distinct band – corresponding to the abundant mRNAs
  3. Store the ds cDNA at –20 until ready to use (within 3 months)
Day 2
Proteinase K Digestion
  1. In a 0.5 ml tube, add:
50 ul of the amplified ds cDNA (2-3 ug)
2 ul proteinase K (20 ug/ul).
  1. Store remaining ds cDNA at –20C for up to 3 months
  2. Mix contents of tube
  3. Spot spin
  4. Incubate 45C, 20 min
  5. Spot spin
  6. Add 50 ul water
  7. Add 100 ul phenol:chloroform:isoamly alcohol
  8. Mix by continuous gentle inversion for 1-2 min
  9. Spin 14,000 rpm for 5 min
  10. Move top (aqueous) layer to a 0.5 ml tube (do NOT pre-chill tube)
  11. Discard interface and lower layers
  12. Add 100 ul chloroform:isoamyl alcohol to the aqueous layer
  13. Mix by continuous gentle inversion for 1-2 min
  14. Spin 14,000 rpm for 5 min
  15. Move the top (aqueous) layer to a 0.5 ml tube
  16. Discard the rest
  17. Add 10 ul 3 M Sodium Acetate
  18. Add 1.3 ul of Glycogen (20 ug/ul)
  19. Add 260 ul of RT 95% EtOH
  20. Spin 14,000 rpm for 20 min at RT
  21. Carefully remove and discard the s/n w/out disturbing the pellet
  22. Wash pellet with 100 ul 80% EtOH
  23. Air dry the pellet (about 10 min)
  24. Add 79 ul water to resuspend the pellet
Sfi I digestion
  1. Combine
79 ul cDNA from above
10 ul 10X Sfi Buffer
10 ul Sfi I enzyme
1 ul 100X BSA
100 ul Total Volume
  1. Mix well
  2. Incubate at 50C for 2 hr
  3. Add 2 ul of 1% xylene cyanol dye to the tube above
  4. Mix well
cDNA Size Fraction by CHROMA SPIN-1000
  1. Invert several times to resuspend gel matrix completely
  2. Hold upright, snap off break-away end (save end)
  3. Put the end into a 2 ml collection tube provided and lift off the top cap (save cap)
  4. Place these tubes into a 13 ml tube with a 1.5 ml tube at the bottom for centrifugation
  5. Spin 2635 rpm (700 x g) for 5 min
  6. Discard collection tube and buffer
  7. Put spin column into a new 2 ml tube or can use 1.5 ml tube for smaller volumes (then put 2 ml tube at the bottom of the 15 ml tube)
  8. Carefully and slowly apply the sample to the center of the gel bed’s flat surface. Do not allow any sample to flow along the inner wall of the column *only load 70-100 ul – can add bromophenol blue to the sample for easier loading (0.01% w/v)
  9. Spin 2635 rpm (700 x g) for 5 min
  10. Remove spin column and collection tube from the rotor and detach them from each other. The purified sample is at the bottom of the collection tube.
  11. Add the following reagents to the tube containing the cDNA: (105-140 ul, respectively)
1/10 vol. Sodium Acetate (3 M; pH 4.8)
1.3 ul Glycogen (20 mg/ml)
2.5 vol. 95% ethanol (-20C)
  1. Mix by gently rocking the tube back and fourth.
  2. Incubate at –20C overnight or place the tube in –20C or dry-ice/ethanol bath for 1 hr (may result in lower recovery)


cDNA Size Fraction by CHROMA SPIN-400
  1. Label 16 1.5 ml tubes and arrange them in a rack in order
  2. Prepare the CHROMA SPIN for drip procedure
    1. Remove spin column from fridge and warm to RT for about 1 hr. Invert column several times to completely resusupend the gel matrix
    2. Remove air bubbles from the column. Use a 1000-ul pippettor to resuspend the matrix gently; avoid air bubbles. Remove bottom cap and let the column drip naturally. – If the column does not drain after 3 min, recap the top cap. This pressure should cause the column to drain)
    3. Attach the column to a ring stand
    4. Let the storage buffer drain through the column by gravity flow until you can see the surface of the gel beads in the column matrix. The top of the column matrix should be at the 1.0 ml mark on the wall of the column. If your column contains significantly less matrix, adjust the volume of the matrix to 1.0 ml mark using matrix from another column.
    5. The flow rate should be about 1 drop/40-60 sec. The volume of 1 drop should be about 40 ul. If the flow rate is too slow (more than 1 drop/100 sec) and the volume of one drop is too small (less than 25 ul), you should resuspend the matrix completely and repeat the drip procedure until it reaches the above parameters.
  3. When the storage buffer stops dripping out, carefully and gently (along the column inner wall) add 700 ul of column buffer to the top of the column an dallow it to drain out
  4. When this buffer stops dripping (about 15-20 min), carefully and evenly apply about 100 ul mixture of Sfi I digested cDNA and xylene cyanol dye to the top-center surface of the matrix. An unsmooth matrix surface does not hurt the following fractionation process
  5. Before proceeding to the next step, allow the sample to be fully absorbed into the surface of the matrix – there should be no liquid remaining above the surface
  6. Wash the tube that contained the cDNA with 100 ul of column buffer by gently applying this material to the surface of the matrix
  7. Let buffer drain until no liquid is left above the resin – at this point the dye layer should be several mm into the column
  8. Place the rack with the collection tubes under the column, so that the first tube is directly under the column outlet.
  9. Add 600 ul column buffer and immediately begin collecting single drop fractions (about 35 ul per tube) in tubes #1-16. Cap each tube after each fraction is collected. Recap the column after fraction #16 has been collected.
  10. Run 3 ul of each fraction on a 1.1% agarose/EtBr gel with a 1 kb marker (0.1 ug). Run at 150 V for 10 min – running the gel longer will make it difficult to see the cDNA bands under UV. Determine the peak fractions by visualizing the intensity of the bands under UV. Collect the first 3 fractions containing cDNA ( in most cases, the 4th fraction containing cDNA is usable. Make sure the fourth fraction matches your desired size distribution). Pool the above fractions in a clean 1.5 ml tube.
  11. Add the following reagents to the tube with 3-4 pooled fractions containing the cDNA: (105-140 ul, respectively)
1/10 vol. Sodium Acetate (3 M; pH 4.8)
1.3 ul Glycogen (20 mg/ml)
2.5 vol. 95% ethanol (-20C)
  1. Mix by gently rocking the tube back and fourth.
  2. Incubate at –20C overnight or place the tube in –20C or dry-ice/ethanol bath for 1 hr (may result in lower recovery)
Day 3
Finish Precipitating out cDNA
  1. Spin 14,000 rpm for 20 min at RT
  2. Remove s/n, do not disturb pellet
  3. Spot spin
  4. Remove remaining liquid and air dry about 10 min
  5. Add 7 ul water to pellet and mix gently.
  6. The Sfi I digested cDNA is now ready to be ligated to the Sfi I digested dephosphorylated lambda TriplEx2 Vector. Start next part or store cDNA at –20C until ready to ligate.

Ligation of ds cDNA to pDNR-LIB
To ensure that you obtain the best possible library from you cDNA, set up 3 parallel ligations using 3 different ratios of cDNA to vector, as shown in Table below. The products of the 3 ligation rxns will be separately transformed into E. coli host cells, and transformations that produce high numbers of clones will be pooled to form the original, unamplified library. In some cases, it may be necessary to perform a fourth ligation using the remaining cDNA in order to obtain a library of the desired complexity.
  1. Label 3 - 0.5 ml tubes and add the indicated reagents (Table below). Mix the reagents gently; avoid producing air bubbles. Spot spin.
Component Ligation A Ligation B Ligation C
cDNA 0.5 ul 1.0 ul 1.5 ul
pDNR-LIB (0.1 ug/ul) 1.0 1.0 1.0
10X Ligation Buffer 0.5 0.5 0.5
ATP (10 mM) 0.5 0.5 0.5
T4 DNA Ligase 0.5 0.5 0.5
Water 2.0 1.5 1.0
Total Volume 5.0 5.0 5.0
  1. Incubate tubes at 16C overnight
Day 4
1. Add 95 ul sterile DEPC-water to each of the ligation rxns.
  1. Add 1.5 ul glycogen
  2. Mix well with pipette tip
  3. Add 280 ul of ice-cold 95% EtOH
  4. Mix by gently rocking back and fourth
  5. Put at –70C or dry ice/EtOH bath for 1-2 hours (Longer incubation times may increase yield of DNA after EtOH precipitation. If transformations are to be performed at a later date, leave the cDNA at –70C.)
  6. Spin 15,000 rpm for 20 min at RT
  7. Remove EtOH layer w/out disturbing pellet
  8. Air dry the pellet
  9. Resuspend each pellet (A, B, C) in 5 ul of sterile DEPC water.
Transformation – Electrocompetent cells
  1. Add 970 ul of LB broth to 14 ml polypropylene tubes
  2. Bring over supplies, samples, etc to Sogin lab
  3. Chill electroporation cuvette for 10-15 mins at –20C
  4. Chill cuvette on ice
  5. Turn machine on by hitting on/standby
  6. To set volts push both arrows at the same time
  7. Hit time const
  8. Thaw electrocompetent cells on ice. Use the cells right after they thaw for maximum efficiency in electroporation. – note: once thawed they can not be refrozen
  9. Add 25 ul thawed cells to each ligation rxn tube ON ICE
  10. Mix thoroughly with pipette tip
  11. Transfer the mixture to a chilled 0.1 cm cuvette
  12. TAP cuvette well and leave cap on
  13. Put into machine quickly
  14. Hit pulse button 2 x (don’t want to hear a pop or else have an arch)
  15. Immediately remove the cuvette from the chamber
  16. Want a reading around 3.5 – 4.5
  17. Add 970 ul LB broth to cuvette and place on ice
  18. Hit on/standby to shut off machine
  19. Transfer the entire volume to the pre-labeled polypropylene tube
  20. Incubate with shaking 225 rpm for 1 hr at 37C
  21. During incubation, add 50 ul LB broth to a 1.5 ml tube and pre-warm small LB/CAM plates (30 min lid off, 30 min lid on)
  22. At the end of incubation, remove 1 ul of the transformation mixture and add it to the 50 ul of LB broth.
  23. Store remaining mixture at 4C
  24. Spread the 51 ul on a prewarmed 90 mm LB/CAM plate
  25. Let soak in for 10 min
  26. Invert and leave overnight at 37C
OR
Transformation – XL10 Gold Kan Ultracompetent Cells (Stratagene #200317 Protocol)
  1. Preheat waterbath to 42C
  2. Pre-chill three (A, B, C) 14 ml BD falcon polypropylene round-bottom tube on ice
  3. Preheat special NZY broth * (w. MgCl2, MgSO4, and glucose) to 42C
  4. Thaw ultracompetent cells on ice.
  5. When thawed, gently mix and aliquote 100 ul cells into each of the pre-chilled tubes
  6. Add 4 ul of the beta-ME mix provided w/ kit
  7. Swirl the tubes gently
  8. Incubate cells on ice for 10 min, gently swirling every 2 min
  9. Add 5 ul of ligation mixture to one aliquot of cells
  10. Swirl tubes gently
  11. Incubate on ice for 30 min
  12. Heat shock at 42C waterbath for 30 seconds
  13. Incubate on ice 2 min
  14. Add 900 ul 42C special NZY broth * (w. MgCl2, MgSO4, and glucose)
  15. Incubate 37C for 1 hr at 225 rpm
  16. During incubation put 50 ul special NZY broth into 3 1.5 ml tubes
  17. Take 1 ul of each transformation mixture and add it to 50 ul special NZY broth
  18. Plate 50 ul of transformation mixture dilution on small (90 mm) LB agar plates with appropriate antibiotic – chloramphenicol @ 30 ug/ml
  19. Allow to soak into plate for 10 min
  20. Store remaining transformation mixture at 4C
  21. Incubate at 37C overnight
Day 5
  1. Determine how many colonies/per are in the library
  2. If high enough to use, store transformation mixture (80%) and glycerol (20%) at –80C in small aliquots (5-10 ul) and use these aliquots to plate out library in the future.
  3. Examine plates.
  4. Pooling transformations for your library
    1. Some of all of the plates should be confluent or nearly confluent. A confluent plate contains several thousand colonies; the corresponding transformation mixture contains 1000 times more independent clones. Three such plates should give you a library of about10^6 clones.
    2. Often one or two of the plates will have considerably fewer colonies than the optimal plate(s). If this is the case (if you do not have 3 confluent plates) we recommend that you perform a 4th ligation using all of you remaining ds cDNA. Choose the ratio of the cDNA to vector based on which of the original ligations A, B, C, gave you the highest number of transformants, and scale up the volumes (see manual). Transform and plate this final ligation according to manual.
    3. If you still do not get 3 confluent or nearly confluent plates, you may wish to repeat the protocol starting with fresh RNA. However, we don recommend that you pool and amplify the clones that you have at this point. These can be combined with the results of your second construction.
    4. (Optional) We recommend that you determine the percentage of recombinant clones in each transformation. To do this, you need to analyze the DNA from 15 independent clones in each transformation. You can screen for inserts by one of two methods: 1) by performing PCR directly on the colonies (see appendix D); or 2) by digestion of miniprep DNA with Sfi to excise inserts. We recommend that you only add a transformation mixture to the pooled library if at least 10 out of 15 clones contain inserts.
    5. Based on the above guidelines, pool the desired transformation mixtures to generate your original, unamplified cDNA library. You need not perform this or subsequent steps with the positive controls.
  5. The original, unamplified library can be stored for up to two weeks at 4C until you are ready to proceed to Amplification of Plasmid Libraries.
Titering Plasmid Libraries
Library Titering Precautions
· Prior to freezing or amplifying your library, you should determine the titer. To ensure a representative library, the titer should be at least 10-fold higher than the number of independent clones. In general plasmid libraries should have a titer of at least 10^8 cfu/ml for long-term storage. Keep the following points in mind when tittering the library
1. Diluted libraries are always less stable than undiluted libraries
2. Once 10^-3 and 10^-6 dilutions of the library are made, use them within the next hour, before drastic reductions in titer can occur.
3. A 2-5 fold range in titer calculations is reasonable, especially if more than one person is doing the tittering.
4. Always use the recommended concentration of antibiotic in the medium to ensure plasmid stability.
5. Use proper sterile technique when aliquoting and handling libraries.
6. Design appropriate controls and include them during plasmid library growth to test for cross-contamination.
Plasmid Library Titering Protocol
1. Prewarm LB/CM plates at 37C (or 30C) for 1-2 hr
2. Thaw an aliquot of the library and place on ice
3. Remove 1 ul of the library, and add it to 1 ml of LB broth in a 1.5 ml microcentrifuge tube. Mix by gentle vortexing. This is Dilution A (1:10^3)
4. Remove 1 ul from Dilution A, and add it to 1 ml of LB broth in a 1.5 ml microcentrifuge tube. Mix by gentle vortexing. This is Dilution B (1:10^6). Note: The diluted library is unstable and should be plated within 1 hr.
5. Add 1 ul from Dilution A to 50 ul of LB broth in a 1.5 ml microcentrifuge tube. Mix by gentle vortexing. Spread the entire mixture onto a prewarmed LB/CM plate
6. Leave plates at RT 15-20 min
7. Invert and incubate at 37C (or 30C) overnight
Day 6
Con’t with Titer
  1. Count the number of colonies to determine the titer (cfu/ml). Calculate the titer according to the following formulas:
1. Colony # Dilution A x 10^3 x 10^3 x 10^3 = cfu/ml
2. (colony # Dilution B/plating volume) x 10^3 x 10^3 x 10^3 = cfu/ml
Amplification of Plasmid Libraries
  • Note: you must amplify the cDNA library to obtain enough high quality plasmid for library screening using ClonCapture and for long-term storage of the library.
Determining the number of Plates Required for Amplification
  1. Note: The exact number of LB/CM agar plates required depends on the library size. Use the following calculation to determine the number of plates to use. Normally, use 3 times the size of the original library and plate at 20,000 cfu/150-mm plate – see manual for example
  2. Calculate the volume of media needed to plate 150 ul on each
  3. Determine what to plate on the plates
Library Amplification Protocol
1. Plate the lib directly on selective medium (LB/CM plates) at a high enough density so that the resulting colonies will be nearly confluent (about 20,000-30,000 cfu per 150 mm plate). Plate enough cfu to obtain at least 2-3X the number of independent clones in the library. (The number of independent clones is the number of independent colonies persent in the library before amplification. To promote even growth of the colonies, continue spreading the inoculum over the agar surface until all visible liquid has been absorbed, and then allow plates to sit at room temp for 15-20 min. If using glass beads to spread the colonies, shake the plate back and forth – not round and round.)
2. Invert the plates and incubate at 37C for 18-20 hr. (Growing the transformants on solid medium instead of in liquid culture minimized uneven amplification of the individual clones.)
Day 7
Storing Amplified Library
1. Add 5 ml of LB + 25% glycerol to each plate and scrape the colonies into the liquid. Pool all the resuspended colonies in one flask and mix thoroughly
a. Set aside five 1-ml aliquots of the library culture in case you wish to re-amplify the library at a later time. Store the aliqotes at –80C
b. Divide the remainder of the library culture into 50-ml aliquots. Use one of the aliquots for ClonCapture as needed. Store the remainder of the library aliquiots at –80C. For use within one week, aliquots may be stored at 4C.



SMART cDNA Synthesis (single)

First Strand cDNA Synthesis
Chill all components and tubes on ice before starting
  1. Combine
1-3 ul RNA Sample (0.025-0.5 ug mRNA or 0.05-1.0 ug total RNA)
1 ul SMART IV Oligonucleotide
1 ul CDS III/3’ PCR Primer
X ul Water up to 5 ul
  1. Mix contents and spot spin
  2. Incubate at 72C for 2 min
  3. Cool on ice 2 min
  4. Spot spin
  5. Add the following
2 ul 5x First-Strand Buffer
1 ul DTT (20 mM)
1 ul dNTP Mix (10 mM)
1 ul PowerScript Reverse Transcriptase
10 ul Total Volume
  1. Mix by gently pipetting
  2. Spot spin
  3. Incubate 42C, 1 hr in an air incubator (says to cover with mineral oil if using thermocycler or waterbath for the incubation)
  4. Put tube on ice to terminate 1st strand rxn
  5. Can stop at this point and leave at –20C if necessary
  6. If continuing on, take 2 ul of 1st strand rxn and put into prechilled 0.5 ml tube
  7. Store remaining 1st strand rxn at –20C for up to 3 months
cDNA Amplification by LD PCR
  1. Preheat thermocycler to 95C
  2. Look at table below to determine how many cycles to run
Total RNA (ug) Poly A+ RNA (ug) Number of Cycles
1.0-2.0 0.5-1.0 18-20
0.5-1.0 0.25-0.5 20-22
0.25-0.5 0.125-0.25 22-24
0.05-0.25 0.025-0.125 24-26
  1. Combine
2 ul First-Strand cDNA
80 ul Water
10 ul 10X Advantage 2 PCR Buffer
2 ul 50X dNTP Mix
2 ul 5’ PCR Primer
2 ul CDS III/3’ PCR Primer
2 ul 50X Advantage 2 Polymerase Mix
100 ul Total Volume
  1. Mix contents by gently flicking tube
  2. Spot spin
  3. Start thermocycling using the following cycle
95C 20 sec
cycle X times (based on table)
95C 5 sec
68C 6 min
  1. In the meantime prepare a 1.1% agarose/EtBr gel
  2. Run 5 ul PCR product with a 1 kb marker – the ds cDNA should appear as a 0.1-4 kb smear with a few distinct band – corresponding to the abundant mRNAs
  3. Store the ds cDNA at –20 until ready to use (within 3 months)
Bacterial Culture Plating – Stock Plates
  1. Streak out XL1-Blue cells onto LB/Tet stock plates
  2. Streak out BM25.8 cells onto LB/Kan/Cam stock plates
  3. Incubate 37C overnight. Can store at 4C for up to 2 weeks
Day 2
Proeinase K Digestion
  1. In a 0.5 ml tube, add:
50 ul of the amplified ds cDNA (2-3 ug)
2 ul proteinase K (20 ug/ul).
  1. Store remaining ds cDNA at –20C for up to 3 months
  2. Mix contents of tube
  3. Spot spin
  4. Incubate 45C, 20 min
  5. Spot spin
  6. Add 50 ul water
  7. Add 100 ul phenol:chloroform:isoamly alcohol
  8. Mix by continuous gentle inversion for 1-2 min
  9. Spin 14,000 rpm for 5 min
  10. Move top (aqueous) layer to a 0.5 ml tube (do NOT pre-chill tube)
  11. Discard interface and lower layers
  12. Add 100 ul chloroform:isoamyl alcohol to the aqueous layer
  13. Mix by continuous gentle inversion for 1-2 min
  14. Spin 14,000 rpm for 5 min
  15. Move the top (aqueous) layer to a 0.5 ml tube
  16. Discard the rest
  17. Add 10 ul 3 M Sodium Acetate
  18. Add 1.3 ul of Glycogen (20 ug/ul)
  19. Add 260 ul of RT 95% EtOH
  20. Spin 14,000 rpm for 20 min at RT
  21. Carefully remove and discard the s/n w/out disturbing the pellet
  22. Wash pellet with 100 ul 80% EtOH
  23. Air dry the pellet (about 10 min)
  24. Add 79 ul water to resuspend the pellet
Sfi I digestion
  1. Combine
79 ul cDNA from above
10 ul 10X Sfi Buffer
10 ul Sfi I enzyme
1 ul 100X BSA
100 ul Total Volume
  1. Mix well
  2. Incubate at 50C for 2 hr
  3. Add 2 ul of 1% xylene cyanol dye to the tube above
  4. Mix well
cDNA Size Fraction by CHROMA SPIN-400
  1. Label 16 1.5 ml tubes and arrange them in a rack in order
  2. Prepare the CHROMA SPIN for drip procedure
    1. Remove spin column from fridge and warm to RT for about 1 hr. Invert column several times to completely resusupend the gel matrix
    2. Remove air bubbles from the column. Use a 1000-ul pippettor to resuspend the matrix gently; avoid air bubbles. Remove bottom cap and let the column drip naturally. – If the column does not drain after 3 min, recap the top cap. This pressure should cause the column to drain)
    3. Attach the column to a ring stand
    4. Let the storage buffer drain through the column by gravity flow until you can see the surface of the gel beads in the column matrix. The top of the column matrix should be at the 1.0 ml mark on the wall of the column. If your column contains significantly less matrix, adjust the volume of the matrix to 1.0 ml mark using matrix from another column.
    5. The flow rate should be about 1 drop/40-60 sec. The volume of 1 drop should be about 40 ul. If the flow rate is too slow (more than 1 drop/100 sec) and the volume of one drop is too small (less than 25 ul), you should resuspend the matrix completely and repeat the drip procedure until it reaches the above parameters.
  3. When the storage buffer stops dripping out, carefully and gently (along the column inner wall) add 700 ul of column buffer to the top of the column an dallow it to drain out
  4. When this buffer stops dripping (about 15-20 min), carefully and evenly apply about 100 ul mixture of Sfi I digested cDNA and xylene cyanol dye to the top-center surface of the matrix. An unsmooth matrix surface does not hurt the following fractionation process
  5. Before proceeding to the next step, allow the sample to be fully absorbed into the surface of the matrix – there should be no liquid remaining above the surface
  6. Wash the tube that contained the cDNA with 100 ul of column buffer by gently applying this material to the surface of the matrix
  7. Let buffer drain until no liquid is left above the resin – at this point the dye layer should be several mm into the column
  8. Place the rack with the collection tubes under the column, so that the first tube is directly under the column outlet.
  9. Add 600 ul column buffer and immediately begin collecting single drop fractions (about 35 ul per tube) in tubes #1-16. Cap each tube after each fraction is collected. Recap the column after fraction #16 has been collected.
  10. Run 3 ul of each fraction on a 1.1% agarose/EtBr gel with a 1 kb marker (0.1 ug). Run at 150 V for 10 min – running the gel longer will make it difficult to see the cDNA bands under UV. Determine the peak fractions by visualizing the intensity of the bands under UV. Collect the first 3 fractions containing cDNA ( in most cases, the 4th fraction containing cDNA is usable. Make sure the fourth fraction matches your desired size distribution). Pool the above fractions in a clean 1.5 ml tube.
  11. Add the following reagents to the tube with 3-4 pooled fractions containing the cDNA: (105-140 ul, respectively)
1/10 vol. Sodium Acetate (3 M; pH 4.8)
1.3 ul Glycogen (20 mg/ml)
2.5 vol. 95% ethanol (-20C)
  1. Mix by gently rocking the tube back and fourth.
  2. Incubate at –20C overnight or place the tube in –20C or dry-ice/ethanol bath for 1 hr (may result in lower recovery)
Bacterial Culture Plating – Working Stock Plates
  1. Pick a single isolated colony form the primary streak plate and streak it onto a LB/MgSO4/Tet (Blue cells) or LB/MgSO4/Kan/Cam plate (BM25.8 cells)
  2. Incubate overnight at 37C. Store at 4C for up to 2 weeks. Use this working stock for inoculating liquid cultures and for preparing the next fresh working stock plate.
Day 3
Finish Precipitating out cDNA
  1. Spin 14,000 rpm for 20 min at RT
  2. Remove s/n, do not disturb pellet
  3. Spot spin
  4. Remove remaining liquid and air dry about 10 min
  5. Add 7 ul water to pellet and mix gently.
  6. The Sfi I digested cDNA is now ready to be ligated to the Sfi I digested dephosphorylated lambda TriplEx2 Vector. Start next part or store cDNA at –20C until ready to ligate.
Ligation of cDNA to lambda TriplEx2 Vector
To ensure that you obtain the best possible library from you cDNA, set up 3 parallel ligations using 3 different ratios of cDNA to vector, as shown in Table below. Use a lambda phage packaging system that will give you at leas 1 x 10^9 pfu/ug of DNA. Follow the supplier’s recommended protocol and perform a parallel packaging rxn with the control insert provided in the packaging kit.
  1. Set up a test ligation to determine the efficiency of ligation the vector to the Control Insert. Use 1 ul of vector, 1 ul Control Insert, 1.5 ul water, and other reagents listed on Table IV. Incubate the test ligation at 16C overnight. Perform a lambda phage packaging rxn and titer the resulting phage. You should obtain >1 x 10^7 pfu/ug of input vector.
  2. Label 3 - 0.5 ml tubes and add the indicated reagents (Table below). Mix the reagents gently; avoid producing air bubbles. Spot spin.
Component 1st ligation 2nd ligation 3rd ligation Test Ligation
cDNA 0.5 ul 1.0 ul 1.5 ul 1.0 control
Vector (500 mg/ul) 1.0 1.0 1.0 1.0
10X Ligation Buffer 0.5 0.5 0.5 0.5
ATP (10 mM) 0.5 0.5 0.5 0.5
Water 2.0 1.5 1.0 1.5
Total Volume 5.0 5.0 5.0 4.5
  1. Incubate tubes at 16C overnight
Inoculate Blue Cells
  1. Prepare 15 ml LB/MgSO4/maltose broth in a 50 ml test tube.
15 ml LB Broth
150 ul 1 M MgSO4
150 ul 20% Maltose
  1. Pick a single, isolated colony from the Blue working stock plate and inoculate broth
  2. Incubate at 37C overnight, 140 rpm until the OD600 is 2.0.

Day 4
Packaging Protocol for Gigapack III Packaging Extract
  1. Perform a separate, lambda phage packaging rxn for each of the ligations
  2. Remove appropriate number of packaging extracts (this number will vary – ask Rick, 2 packages – 2.5 ul/package or 1 package at 4 ul) from the –80C freezer and place the extracts on dry ice
  3. Quickly thaw the packaging extract between your fingers until the extract just begins to thaw (in –80C in gigapack box, blue 1.5 ml centrifuge tube)
  4. Add the sample DNA (ligation mixture) immediately – as soon as ice chunk is gone (1-4 ul containing 0.5 ug of ligated DNA) to the packaging extract
  5. Stir the contents with a pipet tip to mix well. Do not pipet up and down.
  6. Try to avoid air bubbles but if get them it’s okay
  7. Flick tube very gently to mix
  8. Incubate the tube at RT (22C) for 2 hrs. Do not exceed 2 hrs
  9. Note: The highest efficiency occurs between 90 and 120 minutes – so try 100 min
  10. Add 500 ul of SM buffer to the tube (to stop packaging rxn)
  11. Add 20 ul of chloroform (to precipitate protein out)
  12. Mix the contents of the tube gently – mix for 4-5 min, roll tube to disperse chloroform and gently flick continuously for about 5 min, make sure chloroform does not just sit at the bottom – looking for a white precipitate to form
  13. Make sure cap is on tightly
  14. Spin for 2 min at 8000 rpm to collect the protein that precipitated out
  15. Transfer the supernatant to a fresh tube
  16. Store the supernatant at 4C, it is now ready to be tittered
Spin down Blue Cells
  1. Spin cells at 5,000 rpm for 5 min. Pour off the supernatant, and resuspend the pellet in 7.5 ml of 10 mM MgSO4
Prewarm Top Agar and Plates
  1. Warm LB/MgSO4 top agar in microwave
  2. Let cool a bit
  3. Keep in 45C water bath for at least 1 hr before using
  4. Determine the number of small LB/MgSO4 plates you will need for titer
  5. Prewarm plates – lids off 30 min, lids on 30 min
Titering the Unamplified Library
  1. Dilute packaging extracts in 1x Lambda Dilution Buffer. Try 1:5 and 1:20 for an unamplified lambda lysate.
  2. Add 1 ul diluted phage to 200 ul of Blue cells
  3. Put at 37C for 10-15 min
  4. Add 2 ml melted LB/MgSO4 top agar (no hotter than 45C).
*If want to use blue/white selection, add 50 ul IPTG and 50 ul x-gal stock to 2 ml top agar. – Should have about 80% efficiency
  1. Mix by inverting
  2. Pour onto LB/MgSO4 plates and swirl
  3. Let sit 10 min RT
  4. Invert and incubate 37C for 6-18 hr
Inoculate Blue Cells
  1. Prepare 15 ml LB/MgSO4/maltose broth in a 50 ml test tube.
    1. 15 ml LB Broth
    2. 150 ul 1 M MgSO4
    3. 150 ul 20% Maltose
  2. Pick a single, isolated colony from the Blue working stock plate and inoculate broth
  3. Incubate at 37C overnight, 140 rpm until the OD600 is 2.0.
Day 5
Determining the Titer of Unamplified Library
  1. Count plaques
  2. Calc the titer of the phage (pfu/ml)
Pfu/ml=(#plaques x dilution factor x 10^3 ul/ml)/ul of diluted phage plated
  1. If tittering 3 ligations, compare titers to determine the optimal ratio of vector arms to cDNA insert.
  2. From the 3 ligations combined, you should obtain 1-2 x 10^6 independent clones.
  3. Amplify library if titer is good
5. Determine how many plates and what volume to use for amplification. Aim for 6-7 x 10^4 clones (or plaques) per 150 mm plate – thus a library of 1 x 10^6 clones will require 20 plates
  1. The unamplified libraries can be stored at 4C for 2 weeks.
  2. If titer is low religate using the ratio of cDNA to vector that gave the best results but scale up the ligation.
  3. To increase the stability of your library, combine the packaging rxns, and then amplify the library as described below. The amplified library can be stored at 4C for 6-7 months or at –70C (in 7% DMSO) for at least 1 yr.
Prewarm Top Agar and Plates
  1. Warm LB/MgSO4 top agar in microwave
  2. Let cool a bit
  3. Keep in 45C water bath for at least 1 hr before using
  4. Determine the number of large LB/MgSO4 plates you will need for amplification
  5. Prewarm plates – lids off 30 min, lids on 30 min
Spin down Blue Cells
  1. Spin cells at 5,000 rpm for 5 min. Pour off the supernatant, and resuspend the pellet in 7.5 ml of 10 mM MgSO4
Library Amplification
  1. Set up the required number of 4 ml tubes with 500 ul Blue cells and enough diluted lysate to yield 6-7 x 10^4 plaques per 150 mm plate
  2. Incubate in a 37C water bath for 15 min
  3. Add 4.5 ml LB/MgSO4 soft top agar to each tube
  4. Mix, pour onto agar plates, and swirl
  5. Cool 10 min RT
  6. Invert and incubate at 37C for 6-18 hrs, or until the plaques are touching each other
  7. Add 12 ml 1 x lambda dilution buffer to each plate
  8. Store plates at 4C overnight (rotate???)
Inoculate BM25.8 Cells
  1. Pick a single colony from BM25.8 working plate and inoculate 10 ml of LB broth in a 50 ml tube.
  2. Incubate at 31C overnight, 150 rpm, until reach OD600 = 1.1-1.4
Day 6
Preparing Library for Long Term Storage
  1. On a platform shaker (about 50 rpm), incubate the plates at RT for 1 hr
  2. Pour the lambda phage lysates into a sterile beaker
  3. Mix the phage lysate well
  4. Pour it into a sterile 50 ml tube
  5. Add 10 ml of chloroform to the lysate
  6. Screw cap tightly
  7. Vortex for 2 min
  8. Spin at 7,000 rpm for 10 min
  9. Collect s/n to new sterile 50 ml tube cap
  10. Mix gently
  11. Short term storage (up to 6 months)
Aliquot into 1.5 ml screw cap tubes
Store at 4C
  1. Long term storage (up to one year)
Aliquot 1 ml library into 1.5 ml screw cap tubes
Add DMSO to a final conc of 7%
Store at –70C
Avoid repeated freeze/thaw cycles

Day 6 or sometime in the future
Prewarm Top Agar and Plates
  1. Warm LB/MgSO4 top agar in microwave
  2. Let cool a bit
  3. Keep in 45C water bath for at least 1 hr before using
  4. Determine the number of 90 mm (small) LB/MgSO4 plates you will need for amplification
  5. Prewarm plates – lids off 30 min, lids on 30 min
Titering the Amplified Library
  1. Probably could use Blue cells that are one day old or prepare fresh in advance
    1. Prepare 15 ml LB/MgSO4/maltose broth in a 50 ml test tube.
15 ml LB Broth
150 ul 1 M MgSO4
150 ul 20% Maltose
    1. Pick a single, isolated colony from the Blue working stock plate and inoculate broth
    2. Incubate at 37C overnight, 140 rpm until the OD600 is 2.0.
    3. Spin cells at 5,000 rpm for 5 min. Pour off the supernatant, and resuspend the pellet in 7.5 ml of 10 mM MgSO4
  1. Warm 4 LB/MgSO4 (90mm size)
  2. Prepare dilutions of phage lysate – library
    1. Mix 10 ul library into 1 ml 1x lambda dilution buffer (dilution 1 = 1:100)
    2. Transfer 10 ul dilution 1 into a second tube with 1 ml of 1 x lambda dilution buffer (dilution 2 – 1:10,000)
  3. Prepare 4 tubes with 100 ul 1 x Lambda Dilution Buffer, 200 ul Blue cells, and X ul dilution 2: 1 = 5 ul dilution, 2 = 10 ul dilution, 3 = 20 ul dilution, 4 (control) = 0 ul dilution
  4. Incubate tubes in 37C water bath for 15 min
  5. Add 3 ml melted (45C) LB/MgSO4 top agar to each of the four tubes
  6. Mix, pour onto plates, swirl
  7. Leave at RT 10 min
  8. Invert and incubate at 37C for 6-7 hours
  9. Count plaques and calc titer (pfu/ml)
pfu/ml = (# plaques x dilution factor x 10^3 ul/ml)/ul diluted phage plated
Conversion Protocol (Single Excision???)
  1. Add 100 ul of 1 M MgCl2 to the 10 ml overnight culture
  2. Pick a well-isolated positive plaque from secondary or teriary screening plates, and place it in 350 ul 1 x Lambda Dilution Buffer.
  3. Vortex the plaque and incubate at 37C for 3-4 hrs (shake 200-250 rpm???) or allow phage to elute at 4C overnight
  4. In a 20 ml test tube, combine 200 ul of overnight cell culture with 150 ul of eluted positive plaque (save the rest incase need to do again)
  5. Incubate at 31C for 30 min without shaking
  6. Add 400 ul of LB broth
  7. Incubate at 31C for additional 1 hr with shaking (225 rpm)
  8. Using a sterile glass spreader, spread 1-10 ul of infected cell suspension on an LB/carbenicillin plate to obtain isolated colonies and grow at 31C (ampicillin can be used instead of carbenicillin, but may result in more satellite colonies)
  9. Pick colonies, rev prep, sequence
Optional – PCR Insert Screening of SMART PCR cDNA Library
  1. To test for ligation efficiency, screen cDNA insert using Clontech’s advantage 2 PCR Kit and lambda TriplEx LD-Insert Screening Amplimers. Efficient ligation of the cDNA to the lambda TriplEx2 vector should result in more than 80% recombinants.


SMART Reagents

LB Broth

10 g/L Bacto-tryptone
5 g/L Bacto-yeast extract
5 g/L NaCl
(add 15 g agar for plates)
Adjust pH to 7.0 with 5 N NaOH. Autoclave.
LB/Tet Agar Plates - Cool to 50C then add tet (15 ug/ml final conc.)
LB/Kan/Cam Agar Plate – Cool to 50C then add kan (50 ug/ml final conc.) and
Chloramphenicol (34 ug/ml final conc.)
LB Top Agar – 1 L LB Broth, 7.2 g agar. Autoclave.

LB/MgSO4 Broth

1 L LB Broth,
10 ml 1 M MgSO4 (10 mM final conc.). (add 15 g agar for plates) Autoclave.
(1 M MgSO4 – dissolve 24.65 g MgSO4.7H20 in 100 ml water, filter sterilize).
LB/MgSO4/Tet Agar Plates - Cool to 50C then add tet (15 ug/ml final conc.)
LB/MgSO4/Kan/Cam Agar Plates - Cool to 50C then add kan (50 ug/ml final
conc.) and Chloramphenicol (34 ug/ml final conc.)
LB/MgSO4 Soft Top Agar–1 L LB Broth, 10 ml 1 M MgSO4, 7.2 g agar, autoclave.

1 M MgSO4 stock sol’n
Dissolve 24.65 g of MgSO4.7H2O in 100 ml water, filter sterilize.
10 mM MgSO4
Dissolve 2.465 g of MgSO4.7H2O in 1 L water, filter sterilize.
1 M MgCl2
Dissolve 203.3 g of MgCl2 in 1 L water, filter sterilize.

10X Lambda Dilution Buffer

58.3 g NaCl
24.65 g MgSO4.7H2O
350 ml 1 M Tris-HCl (pH 7.5)
Add water to a final volume of 1 L. Autoclave and store at 4C
1 x Lambda Dilution Buffer
100 ml 10X Lambda dilution buffer
5 ml 2% Gelative (0.01% final concentration)
Add water to a final volume of 1 L. Autoclave and store at 4C

Other supplies

1% xylene cyanol dye – 0.1g xylene cyanole ff in 10 ml sterile water = 1% solution
packaging extract
kan stock – 25 mg/ml in water (500x)
tet stock – 15 mg/ml in water (1000x)
chloramphenicol stock – 34 mg/ml in 100% EtOH (1000x)
IPTG 100 mM in water
100% Glycerol
100% DMSO

NZY Broth for use with XL10 Gold Kan Ultracompetent Cells

10 g NZ amine
5 g Yeast Extract
5 g NaCl
Add water to a final volume of 1 L
Adjust pH to 7.5 using NaOH
Autoclave
Add the following filter sterilized supplements prior to use:
12.5 ml 1 M MgCl2
12.5 ml 1 M MgSO4
20 ml 20% (w/v) Glucose (or 10 ml of 2 M glucose)


ZapExpress cDNA Library

Day 1

First Strand Synthesis
  1. Prepare 42C water bath for first strand synthesis
  2. Prepare 16C water bath for second strand synthesis
  3. The final volume of the first-strand synthesis rxn is 50 ul. The volume of added reagents and enzymes is 12.5 ul, thus the mRNA template and DEPC-water should be added in a combined volume of 37.5 ul
  4. Thaw nonezymatic first strand components, vortex, spot spin, keep on ice (vortex and spot spin at 4C the enzyme right before using
  5. In a 0.5 ml RNase-free tube add the following in order (want a total volume of 48.5 ul):
5.0 ul 10 x first strand buffer
3.0 ul first strand methyl nucleotide mixture (called 1st strand dNTP mix)
2.0 ul linker-primer (1.4 ug/ul)
X ul DEPC water (use 0 if mRNA is up to 37.5 ul)
1.0 ul RNase Block Ribonuclease Inhibitor (40 U/ul)
  1. Gently vortex or flick to mix
  2. Add X ul of poly(A) mRNA (5 ug) – total volume should be brought up to 37.5 ul
  3. Gently flick or pipet up and down
  4. Let mixture incubate for 10 min at RT (to let primer anneal to template)
  5. Add 1.5 ul of Stratascript RT (50 ul) – the final volume of the rxn should now be 50 ul
  6. Gently vortex & spot spin
  7. Remove 5 ul of first strand rxn (through away), now you have 45 ul of rxn for second strand synthesis
  8. Incubate first-strand rxns at 42C for 1 hour in a water bath
  9. After incubation spot spin at 4C
  10. Place first-strand rxns on ice
  11. Prepare 72C water bath
Second-Strand Synthesis
  1. Thaw all nonenzymatic second-strand components
  2. Vortex and spot spin tubes
  3. Place on ice
  4. *Make sure all reagents are <16C when the DNA polymerase I is added
  5. To the 45 ul of first-strand rxn on ice, add the following in order:
20.0 ul 10 x second-strand buffer
6.0 ul second-strand nucleotide mixture
116.0 ul good water (DEPC water not required)
  1. Briefly vortex and spot spin enzymes immediately before adding them in the following step
  2. Add the following enzymes:
1.0 ul RNase H (1.5 U/ul)
11.0 ul DNA polymerase I (9.0 U/ul)
  1. Vortex contents of the tube and spot spin at 4C (do quickly)
  2. Incubate for 2.5 hours at 16C (check water bath occasionally to make sure temp does not rise above 16C)
  3. After synthesis for 2.5 hours, immediately place the tubes on ice.
Blunting the cDNA Termini
  1. Add the following to the rxn tube:
23 ul blunting dNTP mix
2.0 ul cloned Pfu DNA polymerase (2.5 U/ul)
  1. Vortex rxn and spot spin
  2. Incubate the rxn at 72C (in a water bath) for 30 min. Do not exceed 30 minutes!
  3. Thaw 3 M sodium acetate
  4. Get about 800 ul of phenol-chloroform and put into a new tube to use later
  5. Remove the rxn, let cool a bit
  6. Add 200 ul of phenol-chloroform [1:1 (v/v)] (don’t use a low pH phenol from the RNA Isolation Kit. Phenol must be equilibrated to a pH of 7-8)
  7. Vortex a lot
  8. Spin the rxn for 2 min at RT at max speed
  9. Transfer the upper aqueous layer, containing the cDNA, to a new tube (0.5 ml) – do not remove any interface that may be present (angle tube and put tip at the top of tube)
  10. Add an equal volume of chloroform
  11. Vortex
  12. Spin the rxn for 2 min at RT at max speed
  13. Transfer the upper aqueous layer, containing the cDNA, to a new tube
  14. Precipitate the rxn by adding the following to the saved aqueous layer:
20 ul of 3 M sodium acetate
400 ul of 100% (v/v) ethanol
  1. Mix by inverting several times or by gently vortexing
  2. Precipitate overnight at –20C
  3. Streak out E. coli XL1-Blue MRF on LB-tetracycling plate
  4. Incubate overnight at 37C

Day 2

  1. Spin max speed at 4C for 1 hr – orient the tube so that can find pellet
  2. The conditions of synthesis and precipitation cause a lg white pellet to accumulate near the bottom of the tube and there may be invisible smaller precipitates, which form along the marked side of the tube. Carefully remove and discard the supernatant.
  3. Gently wash the pellet by adding 500 ul of 70% (v/v) ethanol to the side of the tube away from the precipitate. Do not mix or vortex!
  4. Spin the sample for 2 mins at RT at max speed w/ the same orientation as used before
  5. Remove the ethanol wash
  6. Dry the pellet
  7. Resuspend the pellet in 9.0 ul of EcoR I adapters
  8. Gently flick until dispersed or starts to get dispersed
  9. Incubate at 4C for at least 30 mins to allow the cDNA to resuspend (gently flick every 10 min)
  10. Transfer 8 ul to 0.5 ml tube to proceed in thermal cycler
  11. Save the remaining 1 ul to run on a gel with a 1 kb marker
Ligating the EcoR I Adapters
  1. Add the following components to the tube containing the 8 ul blunted cDNA and the EcoR I adapters:
1.0 ul 10x ligase buffer
1.0 ul 10 mM rATP
1.0 ul T4 DNA ligase (4 U/ul)
  1. Spot spin at 4C
  2. Incubate overnight at 8C in a thermocycler or for 2 days at 4C (optimal ligations occur at 8C). The efficiency of adapter ligation is increased with extended ligation times.

Day 3

  1. In the morning, heat inactivate the ligase by placing the tubes in thermocycler at 70C with heated lid for 30 min
  2. Prepare agarose gel with EtBr and run the 1 ul of 2nd strand rxn on the gel with a 1 kb marker
  3. Prepare SM Buffer is necessary
Phosphorylating the EcoR I Ends
  1. After the ligase is heat inactivated, spin for 2 seconds
  2. Let the rxn cool at RT for 5 min
  3. Separately vortex and spin the following components
  4. Kinase the adapter ends by adding:
1.0 ul 10 x ligase buffer
2.0 ul 10 mM rATP
5.0 ul sterile water
2.0 ul T4 polynucleotide kinase (5 U/ul)
  1. Incubate for 30 mins at 37C in thermocycler (don’t use heated lid)
  2. Heat inactivate the kinase for 30 mins at 70C in thermocycler (use heated lid)
  3. Spin the condensation in the tube for 2 sec
  4. Allow the rxn to come to RT for 5 mins
Xho I Digestion (Cutting XhoI to make it sticky and get rid of GAGA site)
  1. Separately vortex and spin the following components and then add:
28.0 ul of Xho I buffer supplement
3.0 ul of Xho I (40 U/ul)
  1. Incubate for 1.5 hours at 37C in thermocycler (no heated lid)
  2. Cool the rxn to RT
  3. Add 5.0 ul of 10x STE buffer
  4. The sample is now ready to be run though a sephacryl spin columm (do not wait until the next day)
Size Fractionation – Preparing the Column
  1. Get two 1 ml syringes (one for the sample and one for the blank) and two glass1 ml pipets with cotton filter
  2. Get or make sephacryl and invert until homogenous
  3. Take syringe, remove plunger, and remove black stopper off of plunger
  4. Take cotton filter out of pipet and cut in half – keep the bulkier half
  5. Put the cotton in the bottom of the syring using plunger
  6. Put black stopper back on plunger and carefully pack cotton into the bottom of the syringe (be gentle)
  7. Remove plunger
  8. Get 1.5 ml centrifuge tube and cut cap off completely
  9. Get another 1.5 ml tube and cut on the line between the 0.5 and 0.1 lines
  10. Put cut tube under tube with the cap cut off and place in a sterile 15 ml falcon tube
  11. Take syringe and put into 15 ml falcon tube and make sure sits into the centrifuge tube
  12. Make sure sephacryl is homogeneous and suck some up into a pasture pipet (remove any air at the bottom of the tip)
  13. Put mixture into syringe starting at the bottom of the syringe and fill just about to the top (1/4 inch from top). Make sure that you have NO air bubbles. – Do this twice, once for the sample and once for the balance
  14. Spin at 1400 rpm, 20C, for 2 min
  15. Remove flow through and check to make sure plug is working
  16. Add more sephacryl up to about ¼ inch from the top of syringe
  17. Spin at 1400 rpm, 20C, for 2 min
  18. Remove flow through
  19. To wash add 300 ul of 1 x STE buffer (use yellow tips)
  20. Spin at 1400 rpm, 20C, for 2 min
  21. To wash add 300 ul of 1 x STE buffer (use yellow tips)
  22. Spin at 1400 rpm, 20C, for 2 min
  23. Remove collection tube and put a new collection tube in
  24. Pipet the cDNA into the prepared spin column
  25. Spin at 1400 rpm, 20C, for 2 min
  26. Carefully pipet the first fraction (eluent) into a labeled 0.5 ml microcentrifuge tube (Fraction 1)
  27. Replace the column, making sure that the tip is in the microcentrifuge tube
  28. Add 60 ul of 1 x STE buffer on the column
  29. Spin at 1400 rpm, 20C, for 2 min
  30. Carefully transfer the second cDNA fraction to a separate, 0.5 ml labeled tube (Fraction 2)
  31. Add 60 ul of 1 x STE buffer on the column
  32. Spin at 1400 rpm, 20C, for 2 min
  33. Carefully transfer the third cDNA fraction to a separate, 0.5 ml labeled tube (Fraction 3)
  34. You may wish to collect additional fractions, but the size range of the cDNA will decrease.
  35. To the cDNA fractions, add an equal volume of phenol-chloroform [1:1(v/v)]
  36. Vortex
  37. Spin the tube for 2 min at RT at max speed
  38. Transfer the upper aqueous layer to a clean tube
  39. Add an equal volume of chloroform
  40. Vortex
  41. Spin for 2 min at RT at max speed
  42. Transfer the upper aqueous layer to a clean tube – Note: It is very important to phenol-chloroform and chloroform extract the cDNA after the spin column to remove the kinase. Kinase often retains activity after the heat treatment. In the ligation rxn, any remaining kinase activity will allow the vector and the Xho I-EcoR I vector fragment to religate. This may cause a high blue background.
  43. To each extracted sample, add a volume of 100% (v/v) ice cold ethanol that is equal to twice the individual sample volume. Note: The 1 x STE buffer contains sufficient NaCl for precipitation.
  44. Precipitate overnight at –20C

Day 4

  1. Spin the sample for 60 min at max speed at 4C. Transfer the supernatant to another tube. Make sure that almost all of the counts are present in the pellet then discard the supernatant and wash the sample with 200 ul of 80% (v/v) ethanol. DO NOT MIX OR VORTEX! Spin the sample for 2 min at RT and at a max speed. Remove the ethanol and vacuum evaporate for 5 min or until completely dry
  2. Resuspend the cDNA in 4.0 ul of water
  3. Let sit for 10 min at RT
  4. Let sit for 20 min at 4C
  5. Spec with 0.4 ul and 99.6 ul water and determine how many ug/ul you have
  6. Hold at –20C until ready to go to next step
  7. Prepare top agar if necessary (1 L NZY Broth with 7 gm agar)
Ligating cDNA into the ZAP Express Vector Arms
  1. To prepare the sample ligation, add the following components (keep all the components on ICE) and pre-ice tube
X ul resuspended cDNA (about 140-150 ng)
0.5 ul 10x ligase buffer
0.5 ul 10 mM rATP (pH 7.5)
1.0 ul ZAP Express vector (1 ug/ul) (store at 4C not –20C cannot go through another freeze/thaw cycle)
X ul water for a final volume of 4.5 ul
Then add
0.5 ul T4 DNA ligase (4 U/ul)
Then vortex and spot spin at 4C
  1. Incubate the rxn tubes overnight at 12C or for up to 2 days at 4C. The ligated cDNA-vector will form very large concatamers. Breakage of these strands via over-manipulation or multiple freeze-thaw cycles will decrease the efficiency
Preparation of Host Bacteria
  1. Get LB with no antibiotics
  2. Put 20 ml LB into two 50 ml falcon tubes (do under sterile hood)
  3. Mix (separately) by inverting 20% maltose and 1 M magnesium sulfate (both kept at 4C)
  4. Add 200 ul of maltose to 20 ml of LB broth (reverse pipet to avoid contamination)
  5. Add 200 ul of magnesium sulfate to 20 ml of LB broth (reverse pipet)
  6. Get bacteria (XL1-Blue MRF cells that we plated earlier)
  7. Inoculate broth with bacteria
  8. Shake at 200 rpm at 30C overnight

Day 5

  1. Prepare a 47-48C water bath
  2. Pellet the bacteria at 4200 rpm for 10 min in Sorvall – put centrifuge at 4C but can start before it gets down to temp
  3. If pellet is packed, pour off supernatant
  4. Blot and few times and leave upside down for a few min (make sure pellet does not slip)
  5. Gently resuspend the cells in about 5 ml with sterile 10 mM MgSO4
  6. Vortex until cells completely resuspended
  7. Combine the contents of both tubes
  8. Note: the cells may be stored at 4C in 10 mM MgSO4 but should be used within 48 hrs
  9. Dilute the cells to an OD600 of 0.5 with sterile MgSO4
  10. To get cells to 0.5 reading: set up spec at 600 w/ disposable cuvettes
    1. In a fresh falcon tube add about 10 ml 10 mM MgSO4
    2. Add 1 ml stock cell solution
    3. Mix
    4. Take reading
    5. Keep adding 10 mM MgSO4 or stock (reusing the bacteria every time) until get a reading of 0.5, when get reading mix whats in cuvette w/ diluted cells and respec one more time and adjust if necessary (need to come within 0.005)
    6. Keep at 4C
Preparing Top Agar
  1. Microwave and swirl, keep doing until totally in solution
  2. Let cool a little bit before putting into 48C water bath (do not exceed 50C)
  3. Leave in water bath for about 1 hr
Packaging Protocol for Gigapack III Packaging Extract
  1. Remove appropriate number of packaging extracts (this number will vary – ask Rick, 2 packages – 2.5 ul/package or 1 package at 4 ul) from the –80C freezer and place the extracts on dry ice
  2. Quickly thaw the packaging extract between your fingers until the extract just begins to thaw (in –80C in gigapack box, blue 1.5 ml centrifuge tube)
  3. Add the sample DNA (ligation mixture) immediately – as soon as ice chunk is gone (1-4 ul containing 0.5 ug of ligated DNA) to the packaging extract
  4. Stir the contents with a pipet tip to mix well. Do not pipet up and down.
  5. Try to avoid air bubbles but if get them it’s okay
  6. Flick tube very gently to mix
  7. Incubate the tube at RT (22C) for 2 hrs. Do not exceed 2 hrs
  8. Note: The highest efficiency occurs between 90 and 120 minutes – so try 100 min
  9. Add 500 ul of SM buffer to the tube (to stop packaging rxn)
  10. Add 20 ul of chloroform (to precipitate protein out)
  11. Mix the contents of the tube gently – mix for 4-5 min, roll tube to disperse chloroform and gently flick continuously for about 5 min, make sure chloroform does not just sit at the bottom – looking for a white precipitate to form
  12. Make sure cap is on tightly
  13. Spin for 2 min at 8000 rpm to collect the protein that precipitated out
  14. Transfer the supernatant to a fresh tube
  15. Store the supernatant at 4C, it is now ready to be tittered
Plating and Titering
*Do not titer more than 8 at a time*
  1. Warm ‘flat surface’ at 37C
  2. Warm 3 small NZY plates in 37C incubator with lids off
  3. After 30 min put lids back on, and can keep plates in incubator
  4. To plate the packaged ligation product, in a 0.5 ml centrifuge tube make a 1:10 and 1:20 dilution of the product in SM buffer, will also need product that is undiluted
  5. Get 3 2059 tubes and label (S, 1:10. 1:20)
  6. Put 200 ul of XL1-Blue MRF at OD 0.5 all the way to the bottom of the 2059 tubes *make sure cells are homogeneous before pipetting, and vortex or invert often to keep them homogeneous
  7. Put 1 ul of straight sample or dilution right into the cells in the 2059 tube
  8. Jiggle a little bit and get all the fluid at the bottom of the tube
  9. Incubate this (the phage and the bacteria) in an aerated holder at 37C for 15 min to allow the phage to attach to the cells (NO shake)
  10. Every 5 min shake tubes by hand
  11. When done can leave at room temp
  12. Tape the top agar bottle into the waterbath
  13. Add the 3 ml of NZY top agar (48C) to the phage/bacteria mixture
  14. Roll around the tube (about 7 times)
  15. Quickly pour over pre-warmed NZY plate and swirl and shake the plate until totally covered
  16. Remove (or push to the side) any air bubbles using the 2059 tube
  17. When finished with all plates let them sit at RT for 10 min
  18. Put in 37C incubator FACE UP for about 1 hour
  19. Flip up-side-down and leave overnight

Day 6

*Do not work on more than 8 plates at once*
  1. Warm large NZY plates to 37C for at least 1 hour (cover off only 30 min)
  2. Start to liquefy top agar by microwaving, swirl, etc, leave at RT then put into 47-48C water bath
  3. Get plates out and look at which plate you can get the best counts off of. Marker the plate into quarters and count the number of plaques in the four quadrants and total the amount. Determine how many ul of sample you need to get 50,000 pfu but don’t want to exceed 300 ul of phage.
  4. Get 9 2059 tubes into aerated rack and take caps off
  5. Homogenize cell at the OD600 = 0.5 by inverting or gently vortexing
  6. Take 600 ul of cells and put into the bottom of the tubes
  7. Mix plaques gently by inverting or flicking DO NOT vortex
  8. Add plaques that would equal about 10% more than 50,000 pfu/plate (tube) (this time we used 11 ul)
  9. Mix gently by hand
  10. Incubate 15 min at 37C (shake by hand every 5 min)
  11. When done add 7.5 ml top agar
  12. Mix 9-10 times
  13. Pour onto pre-warmed large NZY plates and swirl
  14. Let all sit at RT for 10 min
  15. Record the time finished (we finished at 10:00)
  16. Look at plates every few hours to see if plaques are forming
  17. When plaques are the right size take them out of the incubator (about 6 to 6 ½ hours)
  18. Pre-chill SM buffer to at least 4C and keep as cold as possible when pouring on plates (this is to stop the plaques from growing)
  19. Pour 10 ml of SM buffer onto plates
  20. Swirl around a few times (repeat about 3 times)
  21. Tape up and put on shaker (move it to 4C fridge) on the slowest speed
  22. Grow up XLOLR cell on LB tet plate

Day 7

Preparing Library for Long Term Storage at 4C and –8C
  1. Get plates off of shaker
  2. Wash around (do this quite a bit) and check for dry spots
  3. Pour into 50 ml falcon tubes – do this into 2 tube keeping the volume even
  4. Pour 2 ml SM buffer on each plate
  5. Swirl thoroughly – go once around and then a second time
  6. Pour into the same 50 ml falcon tubes (again splitting evenly)
  7. Mix contents by slowly inverting
  8. Even out the volume of the tubes using a pasture pipet
  9. Estimate the volume (we had 35 ml in each tube)
  10. Add 5% chloroform per total volume (5/95 = X/35 –> so we added 1.8 ml)
  11. Slowly rock back and fourth for a minute or two
  12. Make sure that nothing is leaking
  13. Lay sideways in pink tray with napkins under them
  14. Shake @ RT on #4 for 15 min (periodically check for leaks) – should see white stringy stuff (bacteria and protein) when finished
  15. Spin in Sorvall for 10 min at 7000 rpm (RT/20C)
  16. Take off supernatant with pipet (go from below the surface because probably have some protein at surface that you can see, don’t want to go too close to the chloroform line)
  17. Divide evenly into 4 50 ml falcon tubes
  18. Spin at 7000 rpm for 10 min
  19. Pool all supernatant into a common jar
  20. Mix gently by swirling jar around
  21. To prepare for storage at 4C, in an autoclaved screw cap tube add:
1.5 ml library
5 ul chloroform
invert 10 times to mix, label cap and side, write library down on sheet
  1. To prepare for storage at -8C, in an autoclaved screw cap tube add:
1.5 ml library
113 ul DMSO
5 ul chloroform
invert 10 times to mix, label cap and side, write library down on sheet
Titering Library
  1. Pre-warm top agar
  2. Pre-warm 2 NZY plates (30 min uncovered, 30 min covered) at 37C
  3. Prepare 1.0 OD of XL1-Blue MRF cells
  4. Get tube of library from 4C stock
  5. Make 9 serial dilutions in SM buffer (put 450 ul SM in all 9 tubes, take 50 ul from library and add to dilution 1, vortex, then take 50 ul dilution 1 and put into dilution 2, etc)
  6. Hold dilutions at 4C until ready to use
  7. Get 2 2059 falcons
  8. Put 100 ul of 1.0 OD of XL1-Blue MRF cells into the bottom of the tubes
  9. Add 3 ml of top agar
  10. Mix by inverting about 7 times then pour onto NZY plate, swirl, let cool 10-15 min then put in 4C until ready to use for next step
  11. Grid plates so that they have 9 spaces on them
  12. Drop 10 ul of each dilution onto each plate
  13. Incubate overnight at 37C
Mass Excision
  1. Pre-warm 50 ml NZY broth
  2. Prepare 1.0 OD of XL1-Blue MRF cells
  3. To two 50 ml falcons add then swirl:
375 ul Blue cells at 1.0 OD
1 ul in one tube and 2 ul in the other Library
5 ul Ex Assist helper phage
  1. Incubate 37C for 15 min in aerated holder and swirl every 5 min
  2. Add 20 ml NZY (pre-warmed to 37C)
  3. Incubate at 37C with 200 rpm shake for 2.5 – 3 hours but do not exceed 3 hours
  4. Heat tubes at 70C in water bath for 20 min
  5. Spin 4200 rpm w/ swinging bucket for 10 min
  6. Take off supernatant to sterile tube
  7. Hold at 4C
If plating library out next day streak out a fresh plate of XLOLR cells

Day 7/8

/or sometime in the future

Plating library to use clones for sequencing
  1. Grow up XLOLR – first thing in the morning (8-8:30)
Put 20 ml LB broth (no antibiotics) into 2 50 ml falcon tubes
Add small amt of plated cells
Shake at 39C, 300 rpm until about 3 or 3:30
Spin down (10 min, 4200 rpm swinging bucket, 4C)
Remove supernatant and leave upside-down for a few minutes
Resuspend in 2.5 ml MgSO4, gently vortex to homogenize
Combine contents of both tubes
Get OD600 reading of 1.0
  1. Put 200 ul of 1.0 OD XLOLR cells in the bottom of a 2059 falcon tube
  2. Add 50 ul library (from 4C mass excised stock) directly into cells and gently mix
  3. Shake at 100 rpm at 37C for 15 minutes in aerated holder (push tubes to the bottom of the holder)
  4. Immediately pre-warm small amt of 1 x NZY in falcon tube (only 2-3 mL) to 37C
  5. After 15 minute shake, take out and add 300 ul pre-warmed 1 x NZY (leave at RT when done with NZY)
  6. Put back into shaker at 140 rpm for 45 min (37C)
  7. In the meantime, get appropriate number of lg LB KAN plates and dry at 37C with lids off for ONLY 10 min, then put cover on
  8. When ready to plate always plate a total volume of 100 ul (dilute mixture in the RT NZY if necessary)
  9. Ask Rick how much to plate – sometimes we do 10 ul mix + 90 ul NZY into 1.5 ml tubes
  10. Vortex
  11. Plate out/spread until goes in but don’t be too intense with it


  1. Grow up XLOLR – first thing in the morning (8-8:30)
Put 20 ml LB broth (no antibiotics) into 2 50 ml falcon tubes
Add small amt of plated cells
Shake at 39C, 300 rpm until about 3 or 3:30
Spin down (10 min, 4200 rpm swinging bucket, 4C)
Remove supernatant and leave upside-down for a few minutes
Resuspend in 2.5 ml MgSO4, gently vortex to homogenize
Combine contents of both tubes
Get OD600 reading of 1.0
  1. Put 200 ul of 1.0 OD XLOLR cells in the bottom of a 2059 falcon tube
  2. Add 50 ul library (from 4C mass excised stock) directly into cells and gently mix by hand
  3. Shake at 100 rpm at 37C for 15 minutes in aerated holder
  4. Immediately pre-warm small amt of 1 x NZY in falcon tube (only 2-3 mL) to 37C
  5. After 15 minute shake, take out and add 300 ul pre-warmed 1 x NZY (leave at RT when done with NZY)
  6. Put back into shaker at 140 rpm for 45 min (37C)
  7. In the meantime, get appropriate number of lg LB KAN plates and dry at 37C with lids off for about 10 min, then put cover on (we do not at x-gal to our plates since a high percentage has inserts)
  8. When ready to plate always plate a total volume of 100 ul (dilute mixture in the RT NZY if necessary). Typically a dilution of 20 ul cell mixture + 80 ul NZY works well.
  9. Mix
  10. Plate out/spread until goes in but don’t be too intense with it
  11. Leave plates up-side-down overnight at 37C
  12. Colonies can then be grown up, rev prep’ed, then sequenced



Cod Blood
  1. Spin at 2000 rpm for 10 min.
  2. Separate the red blood cells from the plasma into screw cap tubes.
  3. Store at –20C.



RECENTLY ADDED STUFF




Realtime PCR

Background Lecture on concepts and different types.
Note on price: The brilliant sybr green runs around $160 per 100 rxns so it would be cheaper to use the immomix/syto combo. The syto gets diluted down so much that you can get enough for 250,000 rxns out of a 250ul 5mM bottle.

10 Essential Tips for QPCR [via Bitesize Bio]

Several variations are in use. Needs attention
  • One is with Biolines' Immunomix:
cDNA - X uL
2x Immomix (Bioline) - 12.5uL
10uM Forward Primer - 0.1uL
10uM Reverse Primer - 0.1uL
50uM SYTO13 (Invitrogen) - 1uL
H20 - Add to bring rxn. up to 25uL final vol.

[via Friedman Lab]
...the paper which compares different dyes for real-time. They suggest the optimum concentration of SYTO13 in your recipe to be 2-5uM. I worked out a recipe for a 2uM concentration that seems to be outperforming the brilliant SYBR green kit:
Recipe per 25/ul rxn
12.5 ul of 2X immomix (~$75/100rxns-Bioline)
0.8 ul fwd primer
0.8 ul rev primer
1.5 ul BSA (may exclude this and add volume to water- think it may be causing slight background noise)
1 ul 50uM SYTO13 (~$168 for 250ul of 5mM stock-Invitrogen)
6.4 pcr H2O

Lec16 Realtime PCR
View SlideShare presentation or Upload your own.


Differential Display

via Breeden Lab (FHCRC)



Bleeding Shellfish

Bleeding Clams

[via Jackie DeFaveri]
Other webpage developed for NRAC Field Sampling - sr320 sr320 Nov 24, 2009

Theoretically, we want to hit the heart…..
external image File?id=d8jfhsm_48109wd3vnht_b

But realistically, we are not going to get it, so we actually aim for the posterior adductor muscle
external image File?id=d8jfhsm_4811dd6wgkg4_b

So I hold the clam in the same orientation as shown here, stick the needle in between the shells at about a 45 degree angle. I was using a 21 gauge needle, an inch long, and inserted the whole needle. You should see fluid when you draw back; it should be chunk-free, and a little cloudy. You should easily be able to draw 1ml from animals this size, once the syringe is full you can leave the needle in, remove the syringe to drain in into a tube then re-attach it to the needle and get another ml.
external image File?id=d8jfhsm_4812hc7sjqhh_b


Hope this helps!!



Bleeding Oysters

via Christina Holmes
BLEEDING:
Hemolymph collection from all oysters proceeded the stressing. A notch was made in the shell near the adductor muscle to access the animal. Then the muscle was located by gliding a syringe/needle inside. The needle was inserted into and through the center of the muscle and hemolymph was drawn out. One ml was drawn, the syringe was removed from the needle, and the hemolymph was inspected visually under a microscope to verify the presence of cells. Then a second ml was drawn. Hemolymph samples were held on ice until the cells were separated by centrifugation at 100 x g for 15 minutes. This speed does not break the cells. The supernatant was gently pulled off and transferred to a 2 ml cryovial and was stored with the 5ml falcon tube containing the hemocytes in –80oC freezer. Samples were transported to MA in dry ice overnight then stored at –80C (third shelf down in the bivalve section, in tube racks in blue freezer bags.


Plating Hemocytes

via Rick Goetz
To hemolymph, add additional volume (2 ml seawater/5 ml hemolymph) of cold, sterile-filtered (22 um) seawater containing 100 U/ml penicillin and 100 ug/ml streptomycin (1:100 dilution of stock solution). After gentle mixing, spread 6-7 ml of the hemolymph/seawater solution onto 60 mm culture plates coated with poly-d-lysine. Incubate at 12C O/N to allow for adherence. The following day, GENTLY wash plates with sterile sea water and treat plates as necessary.




Bleeding fish

via Julie Roessig

Here are the supplies we use:
ammonium heparin (Sigma H6279)
18g needles (BD 305196)
1 mL syringe (BD 309602)

You will need 50-100 U of heparin per mL of blood taken. We solublize the heparin in sterile water to a concentration of about 50U/10ul. We then put 10ul aliquots into microcentrifuge tubes and keep them on ice. After drawing the blood, take off the needle and gently squirt the blood into the tube and mix with the heparin before storing on ice. This should keep the blood from clotting without diluting it too much. You can then spin the blood for about 10 minutes at 4 degrees, then remove the plasma and put into a clean tube. Not sure what you are doing after that, but you can store the plasma in the feezer and thaw it later to run certain assays.
==


MSAP

Methylation Sensitive Amplified Polymorphism (MSAP)

protocol here

protocol adapted from:
Utility of the methylation-sensitive amplified polymorphism (MSAP) marker for detection of DNA methylation polymorphism and epigenetic population structure in a wild barley species (//Hordeum brevisubulatum//)
Yidan Li, Xiaohui Shan, Xiaoming Liu, Lanjuan Hu, Wanli Guo and Bao Liu

Polyploid formation in cotton is not accompanied by rapid genomic changes
Liu B, Brubaker CL, Mergeai G, Cronn RC, Wendel JF.



Microbial Community Sampling



Sampling Plan: To sample the microbial communities at various water intake/processing points in conjunction with routine weekly environmental sampling.
General Methods:
Sampling Locations
· Water intake point in the bay
· Post-treated pre-algae water
· Post-treated water + algae
· Larval tank water
Each sample should be 300ml, depending on feasibility of sampling max volume.
Each sample should be taken in duplicate, i.e. 2 samples from each of the 4 locations on each weekly sampling day (n=8 for each day).

Materials needed:
Vacuum manifold - Trap
0.2 um Supor filters (25 mm) [Supor-200 (60300) by Pall or Express Plus (GPWP02500) by Millipore]
tubes for filter paper [Sarstedt, Micro tube 2mL PP No./REF 72.693.005]
rack for tubes
forceps
500 ml grad cylinders
Milli-Q squirt bottle
Preservation solution (0.5M NaCl, 10mM Tris-HCl (pH 8.0), 100mM EDTA (pH 8.0), Filter sterilized!)
Bulb
5 ml pipettes

Filtering the water – Please wear gloves!
  1. Rinse filter funnel with Milli-Q water (distilled and de-ionized) and drain rinse fluid.
  2. Place a 0.2 um Supor filter on filter funnel with tweezers. Carefully put the cups on so that you do not move the filters.
  3. Measure out 300ml of water into a rinsed graduated cylinder.
  4. Pour a little bit of water into the cup to make sure it is sealed. If there are no leaks, fill the cup and begin filtering. Pressure no more than about 10 mmHg. Run through filter.
  5. Aliquot 3mL of preservative (NaCl/Tris/EDTA solution) using plastic pipettor and rubber bulb. Vacuum it out.
  6. Remove the cup. Fold filter in half with the cells on the inside using foreceps.
  7. Place in a labeled tube and then place it in the appropriate box and store at –20 ASAP.
  8. Repeat so we have a duplicate filter for each sampling



DNA Fragmentation (Covaris Sonicator Protocol)

Desired average fragment size = 500bp

Duty Cycle: 5%
Intensity: 3
Cycles per Burst: 200
Time (seconds): 90
Temp (water bath): 4C
Power Mode: Frequency Sweeping
Sample Volume: 120uL
Buffer: TE
DNA Mass: ~8ug
Starting Material Size: >50kb
AFA Intensifier tubes and associated Covaris adapter.


Methylated DNA Immunoprecipitation (MeDIP)

Reagents Needed:
- DNAzol (Molecular Research Center)
- TE
- 100% EtOH
- 70% EtOH
- phenol:chloroform:IAA (25:24:1)
- 5x MeDIP Buffer (50mM Na2HPO4, 700mM NaCl, 0.25% Triton-X 100)
- anti-methyl cytidine antibody (Diagenode; 5-mC monoclonal antibody cl. b)
- Protein A/G Plus Agarose beads (Santa Cruz Biotech)
- 3M sodium acetate (pH = 5.2)
- MeDIP Digestion Buffer (50mM Tris-HCl, pH=8.0, 10mM EDTA, pH=8.0, 0.5% SDS)

Notes:
- Both MeDIP Buffers should be made up from liquid stocks of each individual component, as each individual component are common molecular biology stock reagents that all lab members should have at their benches. Do NOT attempt to make the MeDIP Buffers by weighing out and dissolving powdered chemicals for each individual component; it cannot be done accurately with the small quantities required.

- This is a multi-day procedure. Read through the protocol thoroughly to plan your time properly.


1. Isolate gDNA according to DNAzol protocol (Molecular Research Center), but resuspend final DNA pellet in TE or H2O. (Initial incubation with Proteinase K is dependent on tissue type and available time to perform procedure.)
2. Quantify gDNA yield and quality. Procedure requires a minimum of 6ug, but more can't hurt.

DAY 1

3. Fragment gDNA using Covaris sonicator protocol above. Note: fragmentation protocol requires a sample volume of 120uL.
4. Run 250ng of fragmented on an 2% agarose gel to verify successful fragmentation. Additionally, the fragmentation should be verified/quantified on an Agilent Bioanalyzer if possible.

If fragmentation is successful, proceed to Step 5.

5. Bring fragmented gDNA sample to a volume of 350uL with TE.
6. Heat sample at 95C, 10mins and immediately place on ice for 5mins.
7. Add 100uL of 5x MeDIP Buffer (50mM Na2HPO4, 700mM NaCl, 0.25% Triton-X 100), 45uL of TE and 5uL (5ug) of anti-methyl cytidine antibody (Diagenode; 5-mC monoclonal antibody cl. b). Incubate O/N, 4C rotating end-over-end.

DAY 2

8. Wash calculated volume of Protein A/G Plus Agarose beads (Santa Cruz Biotech; need 20uL per sample) with 1x MeDIP Buffer. Mix stock Protein A/G Plus Agarose beads well and transfer needed volume to clean tube. Pellet beads by spinning 1000g, 2mins, 4C. Discard supernatant. Resuspend beads in 1mL of 1x MeDIP Digestion Buffer. Repeat one time. Final resuspension in 1x MeDIP Buffer. Final resuspension volume is 40uL per sample. Add 40uL of resuspended Protein A/G Plus Agarose beads to each sample and continue incubation with end-over-end rotation @ 4C for 2hrs.
9. Pellet the Protein A/G beads 1000g, 2mins, 4C.
10. Remove and retain supernatant (to retain unmethylated DNA).
11. Wash beads with 1mL 1x MeDIP Digestion Buffer (50mM Tris-HCl, pH=8.0, 10mM EDTA, pH=8.0, 0.5% SDS) by repeating steps 9 & 10. Wash two more times. Save supernatant after each wash.
12. Resuspend beads in 250uL MeDIP Digestion Buffer.
13. Add 70ug of Proteinase K and incubate 24hrs @ RT with end-over-end rotation.
*Note*: The source protocols say to incubate the Proteinase K digest @ 55C. However, we don't have a means to do so, since we need a rotator to keep the agarose beads in suspension. According to various sources, Proteinase K retains >80% of it's enzymatic activity between 20C - 50C. So, allow the digest to run longer (24hrs) than recommended (O/N).

DAY 3

14. Add a volume of phenol:chloroform:IAA (25:24:1) equal to your sample volume, vortex throughly, and spin 16,000g, 10mins, RT.
15. Transfer aqueous phase to clean tube. If original sample had cloudy interphase repeat Step 14 until interphase is no longer cloudy.
16. Precipitate your various DNAs. Add 1/10th volume of 3M NaOAC (sodium acetate; pH = 5.2), 2.5 volumes of 100% EtOH (ethanol), mix throughly and incubate @ -20C for at least 20mins. Note: If expecting low yields, addition of 20ug of glycogen can help improve recovery.
17. Pellet DNA by spinning samples at 16,000g, 20mins, 4C.
18. Discard supernatant and wash pellet with 1000uL 70% EtOH.
19. Re-pellet DNA by spinning samples at 16,000g, 20mins, 4C.
20. Discard supernatant, pulse spin, discard residual supernatant and briefly air dry pellet for 5mins at RT.
21. Resuspend methylated DNA in 50uL TE. Resuspend each unmethylated DNA fraction in 25uL and then pool.
22. Quantify DNAs.

Protocol adapted from:
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2763296/
and
http://www.plosone.org/article/info%3Adoi%2F10.1371%2Fjournal.pone.0013100

5/13/2013 SJW