Phase  

PCR product is run on an agarose gel, the 18s band cut out and cleaned up using a cleanup kit (Zymoclean™ Gel DNA Recovery Kit from Zymo Research). Try to keep the volume of M.Q. (or TE buffer) for elution small to guarantee that the DNA concentration is high enough to establish the desired PCR product: plasmid ratio (see below). Quantify DNA concentration of PCR product (Pico Green^® dsDNA Quantitation Kit from Molecular Probes) and set up Ligation in PCR tubes (pGEM^®-T Easy Vector System I from Promega).

Note: PCR products should be ligated soon after PCR. During PCR Taq adds overhanging A’s on both ends of the 18s fragment. These ends match up with overhanging T’s of the vectors. The A’s tend to fall off after a few days and ligations become far less efficient.

Calculation of PCR product : Plasmid Vector Ratio (3:1 – 1:3)
(see also manual for pGEM^®-T Easy Vector System I)

For 1:1 insert-vector ratio,

50 ng of vector x 1.8 kb size of insert x 1  = 30 ng of insert 
            3.0 kb size of vector               1 

For a 1:1 ratio of DNA product: vector 30 ng of insert DNA are needed. 3 ml of insert DNA can be added to the reaction without exceeding the total preferred ligation mix volume of 10ml (see Table below) => the insert should at least be of a concentration of 10 ng DNA / ml. For a 3:1 ratio a concentration of 30 ng DNA /ml is required.

Example for Ligation Setup:

The ratio of buffer and T4 Ligase (Vector) to total ligation mix volume should be 0.5 and 0.1, respectively. Ratios 1:1 for PCR product to plasmid ratio have worked successfully in the majority of ligation setups.

Note:

• Promega Cloning kits are stored in -80°C. After using the chemicals for the first time they can be stored at -20°C.
• The T4 Ligase and the Vector should be briefly centrifuged (5 sec) before use.
• The 2x Rapid Ligation Buffer should be vortexed vigorously (10 sec) before use.
• Use 20 ml aliquots to avoid temperature changes as the buffer contains ATP.

Ligation takes place overnight at - 4°C in the live fridge.

Prepare SOC Media (needed on DAY 2), S/GAL plates (needed on DAY 2), TB (needed on DAY 3) and clean culture blocks (needed on DAY 3)! Glassware needs to be acid washed (5% HCl) and rinsed with MQ.

*SOC MEDIA  - bacteria are grown in SOC after ligation and before plating. Mix in 1000 ml calibration flask (960 µl of SOC are needed per sample).

20 g                 Bacto tryptone
5 g                   Bacto - yeast extract
10 ml               1M NaCl
2.5 ml              1M KCl

Add tryptone, yeast, NaCl, KCl to ~ 970 ml MQ. Pour into clean 1L Pyrex bottle and autoclave (also autoclave 1 L calibration flask for later use). Let medium cool to room temperature, pour back into autoclaved 1L calibration flask and add

10 ml               2M Mg ²⁺ (stock in live fridge) – 20.33 g MgCl₂.6H₂O plus 24.65 g MgSO₄.7H₂O in 10mls of Q
10 ml               2M Glucose (stock in live fridge) – 36.04g in 100mls Q

using a syringe with a 0.2 mm filter attached (filter sterilize). Bring total volume up to 1000 ml with 0.2 µm filtered, autoclaved MQ. Store media at 4°C (frequently check media for growth).

*TB MEDIA - after picking clones from the SGAL plates they are grown in TB in 96 well culture blocks (Volume needed: 1.25 mls of TB per sample and 120 mls per block).

For 1L TB add:
12 g                 Tryptone                     
24 g                 Yeast Extract                                                 
4 ml                 Glycerol                     
to 900 ml MQ in Pyrex bottle and autoclave

after autoclaving add 100 ml of prepared sterile K solution:

K-Soln for TB Media

Amount of Compound                       Desired Conc.
23.13 g KH₂PO₄                                            0.17 M
125.41 g K₂HPO₄                               0.72 M
up to 1 liter with Milli-Q, Autoclave 0.17 M

AND when solution is < 50°C (tolerable to touch) add 1ml of 100 mg/ml Ampicillin
Media is stored at 4°C.

* S/GAL plates for growth of colonies (needed on DAY 2). 2 packages (Premix from Sigma Aldrich C-4478) is dissolved in 1L MQ and autoclaved (500 ml of media allows to pour appr. 30 plates). Let the medium cool down “tolerable to touch” (30-45 min under live hood) and add 1ml of Ampicillin (100 mg/ml stock) to 1 L of medium. The medium is swirled and poured into large petri dishes avoiding bubbles. After the plates solidify they are put back in the plastic sleeve and stored in the live fridge. Turn plates upside down to avoid condensation and label them (eg S/Gal + Amp, date).

* Culture blocks (96 wells, 2ml Polypropylene from Whatman®) are needed on DAY 3 to grow up bacteria. Should be acid washed, MQ rinsed and autoclaved.

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Phase  

Set incubation chamber/shacker to 37°C.

LIGATION CLEANUP

• After ligations have sat overnight, use “StrataClean Resin” to clean up the reaction (provided with Electro Ten-Blue^® Electroporation Competent Cells from Stratagene).
• Vortex the StrataClean resin to completely resuspend it.
• Pipet 5µl of the slurry into each tube containing the ligation reaction.
• Vortex this mixture for 15 sec.
• Pellet the resin by centrifugation at >2000xg for 1min.
• Carefully remove the supernatant containing the DNA to a fresh microcentrifuge tube. !!Remove only a maximum of 10 µl of supernatant!!
• Repeat steps 1-5
• Use the supernatants in the following electroporation reaction.  Aim for 5ng DNA to be added to 40 µl of electro-competent cells (for best ligation results PICO quantification after Resin cleanup is suggested).

ELECRTOPORATION (Biorad Gene Pulser Xcell system)

Have ready: Electroporation cuvettes and 1.7 ml microcentrifuge tubes chilled on ice and SOC media (960µl per sample) preheated to 37°C.

• Thaw the electroporation competent cells (Electro Ten-Blue^® Electroporation Competent Cells from Stratagene) on ice (~ 5 min).
• For each ligation sample aliquot 40 µl of electrocompetent cells into chilled 1.7 ml microcentrifuge tubes.
  *Electrocompetent cells are very fragile, handle with care when pippeting.
• Add experimental DNA (~ 5ng plasmid DNA) to 40 µl electroporation-competent cells. Gently mix the cells and DNA.
  Positive Control: Dilute the pUC18 control plasmid (provided with electrocompetent cells) 1:10 with sterile dH₂O.  Add 1 ?l of the diluted pUC18 control plasmid to 40 µl of cells.
• Transfer each DNA-cell mixture to a chilled electroporation cuvette. Tap the cuvette until the mixture settles evenly to the bottom!!
• Wipe the sides of the cuvette to remove any water and slide the cuvette into the chilled electroporation chamber until the cuvette connects with the electrical contacts.
• Shock the sample and quickly add 960 µl of the 37°C sterile SOC media to resuspend the cells (electroporation conditions see Manual from Stratagene).
• Transfer each cell suspension to a sterile 15ml Falcon polypropylene tube and incubate at 37°C for 90min shaking at 225-250 rpm.
• Plate ~100-200 ml of cell suspension onto duplicate/triplicate S/GAL plates using wheel table and “looped sticks”. Between every sample dunk loop into 90% Ethanol and fire sterilize. Make sure loop cooled down enough by touching S/GAL on the outer rim. Leave plates under hood for a few min (~ 5 min) so the streaked cell suspension dries.
• Incubate plates upside down at 37°C for ~24 hrs.

* Make sure glasware and pipettes are exposed as Biohazard after they come in contact with electrocompetent cells.

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Have ready: Cleaned culture blocks (HCl washed, MQ rinsed and autoclaved), autoclaved toothpicks. When picking clones work in live hood to avoid contamination!

• Fill 1.25 ml of TB media into all 96 wells of the culture block.
• Use autoclaved toothpick to transfer cells from single colonies on the S/GAL plate to corresponding wells filled with TB media (toothpicks can be left in the wells or just immersed and removed). Inoculate one colony in each well. Keep track of sample IDs and additional information.
• Seal the wells with “Airpore Tape sheets” (Qiagen) and incubate blocks for 16-24 hours at 37°C and ~ 300 rpm. Use tape to fix wells onto bottom of shaker. (Shaking the blocks at more than 300 rpm causes cross contamination!)
• Clean hood using 90 % Ethanol. Dump S/GAL plates, toothpicks, etc… into Biohazard trash.

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Remove blocks from shaker, spin them briefly and carefully take of Airpore tape (potential for contamination if drops of sample are stuck to Airpore tape). Load 85 ml of cell suspension from each well into a 0.7 ml microcentrifuge tube preloaded with 15 ml Glycerol (should be ~15% of total volume) and vortex.

These samples serve as BACKUPS for the clone library and are stored at -80°C.

To regrow samples: thaw, transfer 5-10 µl of each cell mix into culture blocks or tubes holding fresh TB media (see Cloning Day 3).

Close the blocks after removing cells for creating backup libraries with fresh adhesive tape and centrifuge at 20°C at 1913 x g for 30 min to spin down the cells. Remove tape carefully and then remove media by inverting blocks. Do not agitating the block too much as the cell pellet might detach and be lost.

• Resuspend pellet by adding 250 ml Cell Resuspension Solution to each well (if the block was stored frozen prior plasmid purification, make sure the pellets are completely thawed). Letting the pellets sit in the cell suspension for 5-15 minutes might increase recovery. Thoroughly mix until a uniform cell suspension is achieved (mix by pipetting samples up and down repeatedly or by agitating the whole block using a vortex mixer).
• Add 250ml of Cell Lysis Solution to each sample and mix by tapping plate against the palm of your hand 3-4 times. Incubate at room temperature for 3 min.
• During incubation place DNA Binding Plate (blue dot in top left-hand corner of plate) on Manifold Base and connect vacuum port to vacuum source using the insert provided with Manifold and vacuum tubing. Place Manifold Collar on top of the base and binding plate by aligning the collar with the pins. Make sure numbering and orientation are lined up correctly.
• Add 350 ml of Neutralization Solution to each sample (no mixing necessary). Transfer bacterial lysates to the Lysate Clearing Plate assembled on the Vacuum Manifold. Make sure the tips are disposed into the Biohazard trash. Allow filtration disks to wet uniformly for 1 min before applying vacuum (15-20 inches Hg). Allow Lysate to pass through clearing and DNA-binding plate (3-5 min). All lysate should have passed through both plates. Release vacuum. Remove Clearing plate and collar. Depending on the thickness of the cell suspension this step might take longer. Dispose of Lysate plate.
• Add 500ml of Neutralization Solution to each well of the DNA Binding plate, apply vacuum for 1 min and switch off pump.

• Add 1ml of Wash Solution containing EtOH (95% EtOH is to be added to the washing solution as indicated on the bottle) to each well of the DNA-binding plate. Apply vacuum for 1 min.
• Repeat wash procedure! Continue vacuum for another 10 min after wells have emptied to allow binding matrix to dry. Release vacuum line from Manifold Base and snap it into the vacuum port in the Vacuum Manifold Collar. Remove Binding plate from Manifold Base and blot by tapping onto a clean paper towel to remove residual EtOH.
• Place a 96 well Elution Plate in the Manifold Bed and position the Vacuum Manifold Collar on top. Make sure numbering and orientation are lined up correctly.

• Position Binding-Plate on top of Manifold Collar and Elution Plate. Make sure that Binding-Plate tips are centered over the Elution Plate wells. Add 100ml of Nuclease-free Water (make sure water is DEPC-free, might inhibit sequence reactions) to each well of the Binding Plate and incubate for 1-2 min at room temperature. Apply vacuum for 1 min.
• Release vacuum, tap the Binding plate forcefully to make sure that all liquid containing DNA is caught in the elution plate wells and remove Binding Plate. Carefully remove Manifold collar making sure that the Elution Plate remains positioned in Manifold bed. If any droplets get onto the walls of the wells blocks, the plate should be briefly centrifuged so that elute is at bottom of well. Normal recovery is about 60-70ml.

Alternatively elute plasmid DNA by centrifugation: the DNA Binding-plate and elution plate are assembled like described above and centrifuged at  ~ 4000 rpm for 1 min. This has generally resulted in better recovery of plasmid DNA.

Plasmids can be used directly for sequence reactions or should be stored at -20°C.

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