Lab 7 (Modified): Subcloning, ligation and transformation

 

Note the shortage of competent cells is not as severe as was reported

 

In this lab, a 5.3 kb PstI fragment will be purified from an agarose gel, subcloned into PstI-cut pUC19, and transformed into DH5α (DH5alpha) competent cells.

 

Elution of gel band

 

Before the lab, plasmid pPBH (pBluescript containing a fragment of P. ochraceus histone gene sequence) will be digested with PstI. 0.5 g aliquots of the digested DNA will be loaded on agarose gels, along with a 1 Kb Ladder molecular weight marker. After the gels have run long enough for the bromophenol blue band to be near the end of the gel, two bands should be visible in the sample lanes: a 5.3 kb histone gene fragment, and a 3 kb vector fragment. The 5.3 kb histone gene band will be cut out of the gel, and eluted using the GFX PCR DNA and Gel Band Purification Kit. This is a kit which uses NaI to dissolve agarose and a silica matrix column to bind the eluted DNA. For instructions on how to use the kit, see the Subcloning lab references on reserve in the library. The most important step in using the kit properly is to cut out the gel band in the minimum amount of agarose possible. This will reduce the amount of the capture buffer needed to completely digest the agarose. Also be careful not to expose the DNA to UV light for more time than is necessary this will minimize DNA damage due to UV radiation.

 

Assuming complete recovery of the band, the amount of DNA recovered should be a fraction of the 0.5 g of DNA loaded on the gel proportional to the percent that the 5.3 kb fragment made up of the original 8.3 kb plasmid. (E.g. 0.5 g X 5.3 kb/8.3 kb). The fragment will be eluted from the columns using 50 l sterile ddH2O. (TE could also be used for this purpose, but the EDTA might interfere with the ligation step. T4 DNA ligase requires Mg2+ as a cofactor.)

 

Protocol:

 

1. When bromophenol blue band is near end of gel, remove gel from apparatus and place on saran wrap. Photograph the gel, then take it to the darkroom in lab 8159.

 

2. Follow the protocol for the GFX kit (see the Subcloning lab references on reserve in the library).

 

3. Elute the DNA using 50 l sterile ddH2O. Assuming complete recovery of the band, calculate the concentration of the DNA present.

 

Ligation

 

For maximum yield, ligations are usually done at 14-16C overnight. (1-4 hours at room temperature will also give decent yields.) Because of time constraints, the ligation in this lab will be done using a Rapid Ligation protocol suggested by Invitrogen/Life Technologies, which is supposed to produce results in 5 minutes at room temperature. The main difference between this and the standard ligation protocol is the higher concentration of ligase used 1 Weiss unit per reaction, instead of the 0.1 unit per reaction that is normally used for cohesive end ligations. (See the Subcloning lab references on reserve in the library). To increase the chances of ligation between vector and insert, a ratio of 3 insert DNA ends to 1 vector DNA end will be used. Since ligase joins DNA ends together, the concentrations of DNA ends (in fmols) for both DNA molecules must be known.

 

fmols DNA ends can be calculated using the following formula:

 

1 g DNA (for a 1 kb fragment) = 3000 fmol DNA ends

 

g DNA = fmol DNA ends X 1 g/3000 fmol X size of DNA in kb/1 kb

 

(See the Subcloning lab references on reserve in the library for examples of calculations).

 

Two ligations will be set up by each group a test ligation containing insert and vector DNA, and a control ligation containing only vector DNA. Because T4 DNA ligase is not stable for long at 0C, the 1 unit/l ligase working stock will be prepared each day shortly before use. T4 DNA ligase buffer contains ATP, so it must be kept on ice while being used. The polyethylene glycol (PEG) in Life Technologies DNA ligase buffer may precipitate during freezing therefore thawed buffer should be vortexed vigorously before use.

 

Materials

 

1 unit/l T4 DNA ligase (in 1X Life Technologies DNA ligase buffer) working stock

5X DNA ligase buffer

eluted 5.3 kb PstI fragment

PstI digested pUC19 (6.5 ng/l) (heated to 80C for 20 minutes to inactivate enzyme)

sterile ddH2O

 

Methods

 

1. Each group will prepare the following ligation mixes:

Ingredient

Test ligation

Control ligation

5X DNA ligase buffer

4 l

4 l

vector DNA

14 fmol DNA ends

14 fmol DNA ends

insert DNA

42 fmol DNA ends

none

sterile ddH2O

to 20 l

to 20 l

1 unit/l T4 DNA ligase

1 l

1 l

Total volume

20 l

20 l

 

2. Pulse spin the tubes to concentrate the ingredients, mix by flicking with finger.

3. Incubate at room temperature for at least 5 minutes.

 

4. Dilute to 100 l total volume with sterile ddH2O before transforming competent cells. (This will improve transformation efficiency.)

 

Chemical Transformation of E. coli

 

The ligation mixes will be transformed into commercially prepared chemically competent E. coli DH5α.

 

DH5α genotype:

supE44 deltalacU169 (phi80 lacZ deltaM15) hsdR17 recA1 endA1 gyrA96 thi-1 relA1

 

If you wish to see recipes for making competent cells, consult the Sambrook reference in the lab.

 

Each group will do 5 transformations: the test ligation and 4 controls. The controls will be the control ligation, unligated PstI-cut pUC19 vector DNA, pPBH plasmid miniprep (from Lab 3), and a mock transformation with no DNA added. Different amounts of DNA are needed for the different reactions, because supercoiled plasmid DNA transforms competent E. coli cells more efficiently than open-circle ligated circular plasmid DNA. Linear DNA transforms chemically competent E. coli at a very low rate.

 

Materials (per group):

 

7 plates (LB + 100 g/ml ampicillin)

X-gal (20 mg/ml)

IPTG (20% w/v)

1 X 125 l aliquot of competent cells

diluted test ligation (100 l)

diluted control ligation (100 l)

unligated PstI-cut pUC19

pPBH plasmid miniprep (from Lab 3)

 

1. Aliquot 1 X 125 l aliquot of competent cells from one of the stock tubes. Divide this into 5 X 25 l of competent cells on ice in 5 tubes.

 

2. Set up the following transformations. Tubes should be kept on ice while you set up the mixes.

 

Ingredient

Test ligation

Control ligation

Unligated digested pUC19

pPBH plasmid miniprep

Mock ligation

Competent cells

25 l

25 l

25 l

25 l

25 l

DNA

10

10 l

2 l

~0.5 l

none

 

3. Leave transformation mixes on ice for 10 minutes.

 

4. Swirl the tubes in the 37C waterbath for 2-3 minutes. Return to ice.

 

5. Add 900 l of LB and incubate in the 37C waterbath for 1 hour.

 

6. Prepare LB + ampicillin plates by adding 40 l of X-gal and 4 l of IPTG. Spread evenly over plate using a sterile hockey stick. Label the plates properly.

 

7. Centrifuge the transformation mixes for 30 seconds, remove the supernatant, and resuspend the pellet in 50 l of LB.

 

8. Plate 40 l of test ligation transformants on one plate, and 10 l on a second plate.

 

9. Repeat step 8 for the control ligation.

 

10. Plate 50 l of the unligated digested vector and mock transformations.

 

11. Plate 15 l of the pPBH plasmid transformation.

 

12. Incubate overnight at 37C and count blue and white colonies the next day.

 

Study questions (due next week)

 

1. What were the purposes of the various control transformations used?

 

2. What percent of the colonies on your test ligation plates contained cloned inserts?

 

3. Calculate the yield of transformants per ng of DNA for the test ligations.

 

4. Why were DH5alpha competent cells used for the transformations? What gene makes this possible?