Unless stated on the top of each page, all protocols were adapted and modified from Campbell (1998).
 

A. Polymerase Chain Reaction (PCR)

PCR is used to amplify a specified DNA sequence. Oligonucleotide primers are used to flank the region of the DNA to be amplified. Repeated cycles of heating and reannealing allow exponential synthesis of the desired DNA fragment.

The reagents needed for PCR include:
* Template DNA
* Reaction buffer with or without 15 mM Mg²⁺ (10 X stock solution)
* MgCl₂  (25 mM)
* dNTPs (20X or 4mM stock solution)
* Forward primer (100X stock solution)
* Reverse primer (100X stock solution)
* Taq DNA polymerase (5 u/ml)

Although PCR is traditionally done in 100 ml aliquots, all reactions in my experiment were carried out in 50 ml volumes. Appropriate concentration of Mg²⁺ ions  is crucial in PCR success. Using the correct amount of Mg²⁺ can ensure maximal PCR efficiency.

REAGENTS                                                    VOLUME (ml)
dH₂O                                                                     50-X
Template DNA                                                          X
Reaction buffer (w/ or w/o 15 mM Mg²⁺ )                 5.0
MgCl₂ (25 mM)                                                       2.0
dNTPs (20X or 4 mM)                                              2.5
Forward primer (200 ng/ml)                                      0.5
Reverse primer (200 ng/ml)                                       0.5
Taq DNA polymerase (5 u/ml)                                  0.5
Total                                                                      50.0

Once all reagents are mixed in a 500 ml microfuge tube, the mixture was placed in the PCR thermal cycler and the GA1 program was run.

GA1 Program
Step 1 : 95.0° C for 5 minutes
Step 2 : 95.0° C for 1 minute
Step 3 : 55.0° C for 1 minute
Step 4 : 72.0° C for 2 minutes
Step 5 : Repeat Steps 2-4 for another 29 cycles
Step 6 : Maintain at 20.0° C

B. "Cleaning"  DNA

"Cleaning" of DNA samples refers to the removal of proteins and salts from the solution. DNA is  often "cleaned" after restriction enzyme digestions or PCR. If the digestions are to be further processed or manipulated. This "cleaning" process involves precipitation and extraction of the DNA using ethanol.

1. The volume of DNA in the tube was brought to 200 ml with dH₂O if the total volume was less than 200 ml.
2. An equal amount of phenol/chloroform/isoamyl alcohol was added to the solution, vortexed for 30 seconds and  then centrifuged for 2 minutes at 12,000 rpm.
3. The upper aqueous layer that contains the DNA was then pipetted out and placed in a new microfuge tube.
4. The entire extraction procedure was repeated with 100% chloroform.
5. 20 ml of 3M sodium acetate and 800 ml of 4° C 100% ethanol was added to the solution, vortexed for 30 seconds before putting the tube in a -20° C freezer overnight or in the -70° C freezer for 30 minutes.
6. The tube was then centrifuged at 12,000 rpm for 10 minutes.
7. The supernatant was removed by aspiration.
8. The pellet at the bottom of the tube was washed with 500 ml of 4° C 70% ethanol and dried by the Savant DNA 110 Speed Vac^TM  .
9. The pellet was resuspended in appropriate volume of TE (Tris-EDTA) buffer.
 

TE Buffer- Tris, pH 8.0, EDTA
To make 50.0 ml
5 ml of 1M Tris, pH 8.0
1 ml of 0.5 M EDTA
44 ml of H2O

A typical 10 ml ligation reaction consists of the digested vector, targeted insert, sterile water, ligation buffer and T4 DNA ligase. The ligation reagents were mixed in a 500 ml microfuge tube, and the tube was incubated in a thermal cycler at 16° C for 4 hours with the heated lid disabled.

Ligation reaction table :

Digested Vector (50 ng)                                 1.0 µl
*Insert (2:1 molar ratio insert:vector)                x µl
Sterile water                                          7.5 µl- x µl
10X ligation buffer                                        1.0 µl
T4 DNA ligase (keep cold)                            0.5 µl
Total Volume                                               10.0 µl

Calculation for amount of insert used at a 2:1 insert to vector ratio :

X ng of insert = (2) (bp insert) (50 ng linearized plasmid-) ÷ (size of plasmid in bp)

* The insert to vector ratio can be easily changed to 4:1, 8:1 or 10:1 depending  on the transformation reaction being carried out. The amount of insert to be used can be calculated from the formula by substituting the 2x multiplier with the desired ratio of insert used. In many of my experiments, a 4:1 or 8:1 insert to vector ratio was favored since the chances of getting more ligated inserts is higher.

1. A solution of 100 mM glucose in a 1% agarose gel was prepared.
2. 1.5 mL of the molten agarose was poured into as many 1.5 mL microfuge tubes as needed.
3. A piece of tape with a small hole in the center was used to cover the top of each tube so that the hole resided in the middle of the tube opening.
4. A yellow pipet tip was then pushed through the hole into the molten agarose  to a depth of  60 ml. Gel was allowed to solidify.
5. Once the molten agarose gelled, the pipet tip and the tape was removed from the tube opening.
6. The ligation reaction was carefully pipeted into the agarose well.
7. The tube was then allowed to sit on ice for 90 minutes to allow diffusion of ligation salts.
8. The ligation mixture was  removed from the well and electroporation was then carried out.

I) Heat-shock transformation

1. Chemically competent E. coli cells were used in all the heat shocked transformations.
2. The competent E. coli strains used in the transformations :
    a) JM109 High Efficiency Competent Cells from PromegaÇ
    b) MAX Efficiency STBL2 Competent Cells from Gibco BRL Ç
3. Prior to transformation, the microfuge tubes to be used were chilled in an ice bucket.
4. 5 ml of the  ligation reaction was then pipetted into the chilled microfuge tubes.
5. The competent cells were then obtained from the -70° C freezer and thawed for 7 minutes. 40 ml of the thawed cells were then added into the chilled microfuge tubes which contains the ligation reaction
6. The DNA and cells were then placed on ice for 10 minutes (JM109) or 30 minutes (STBL2).
7. The cells were then heat shocked in 42° C heat bath for 2 minutes (JM109) or 25 seconds (STBL2).
8. If STBL2 cells were used, the tube was then placed on ice for an additional 2 minutes.
9. 1 mL of Luria Broth (LB) medium was then added into each tube and the cells were allowed to recover in a 37° C incubator for 1 hour.
10. Cells were then centrifuged at 12,000 rpm for 20 seconds and the supernatant was discarded.
11. The pellet was then dissolved in fresh 100 ml LB medium and plated on the an appropriate LB + antibiotic plate. Plates were dried and incubated overnight at 37° C.

II) Electroporation transformation

1. Epicurian Coli SURE electroporation competent cells from StrategeneÇ were used in all electroporation transformations.
2. The electrocompetent cells were thawed for 7 minutes prior to transformation.
3. 5 ml of the ligation reaction were then mixed with 40 ml of the electrocompetent cells in a pre-chilled microfuge tube.
4. Using the provided dropper, the mixture was transferred from the tube into a pre-chilled electroporation cuvette.
5. The electroporator was set at 1.3kV and the cuvette, wiped dried from any moisture at the surface, was placed within the electrode chamber.
6. The charge was then released and a pulse was fired.
7. 960 mL of LB was added into the cuvette and the electroporated cells were then transferred to a sterile microfuge tube. Cells were then allowed to recover for 1 hour at 37* C in an incubator that was shaking at 225 rpm.
8. The recovered cells were centrifuged for 20 seconds, the supernatant was discarded and 100 ml of fresh LB was used to resuspend the pellet.
9. The 100 ml of resuspended cells were then plated on the appropriate LB + antibiotic plate.
10. The plate was incubated overnight at 37° C.

F. Alkaline Lysis Mini-Prep

1. For each individual mini-prep, 3 mL of  the cell culture was grown overnight in an appropriate LB + antibiotic medium at 37° C and shaken at 400 rpm.
2. The contents of the cell culture was then poured into a 1.5 mL microfuge tube and the remainder was stored in a fridge.
3. The microfuge tube was then centrifuged for 2 minutes at 12,000 rpm and the supernatant was aspirated.
4. The pellet was resuspended in 100 ml of Mini-Prep solution I.
5. 200 ml of Mini-Prep solution II was then added and the mixture was vigorously shaken.
6. 150 ml of ice cold Mini-Prep solution III was added and the solution was gently mixed.
7. The microfuge tube was then centrifuged for 5 minutes at 12,000 rpm and the supernatant was transferred into a fresh microfuge tube.
8. Extraction with 400 ml of phenol/chloroform/isoamyl alcohol was carried out. Tube was vortex and centrifuged for 2 minutes at 12,000 rpm.
9. The upper aqueous layer was extracted into a fresh microfuge tube.
10. The entire extraction process was repeated with 100% chloroform.
11. 800 mL of room temperature 100% ethanol was added to the supernatant from the second extraction. The mixture was vortexed and left to stand for 2 minutes at room temperature.
12. The tube was then centrifuged for 5 minutes at 12,000 rpm at room temperature. The supernatant was then discarded and the pellet was washed with 500 ml of ice cold 70% ethanol.
13. The pellet was then dried with a Savant DNA 110 Speed Vac^TM.
14. The pellet was then dissolved in 50ml of TE buffer and 5 ml of RNase A.

Mini-Prep Solution I
50 mM glucose
25 mM Tris, pH 8.0
10 mM EDTA, pH 8.0

Mini-Prep Solution II
0.2N NaOH
1% SDS
0.4 ml of 5 M NaOH
0.5 ml of 20% SDS
9.1 ml of dH2O (SDS and NaOH are not mixed directly, water added first)

Mini-Prep Solution III
3 M K and 5 M acetate
for 100 ml:
60 ml 5 M K Ac
11.5 ml glacial acetic acid
28.5 ml dH₂O

G. Restriction Digestion

A restriction enzyme is capable of cleaving  a specific double-stranded DNA sequence. To carry out a restriction digest, optimal conditions have to be met to increase the efficiency of the enzyme. Optimal conditions include having the correct temperature, salt concentrations and optimal pH. This means using the correct buffer is crucial in a restriction digest.  In addition, since the restriction enzyme is stored in glycerol, the appropriate amount of enzyme has to be used to prevent star activity (<5% v/v).

Restriction digest :

  5 µl    DNA (volume depends on DNA concentration, 5 µl for mini-prepped  DNA)
12 µl    water (up to desired volume)
  2 µl    10X buffer (one tenth final volume)
  1 µl    restriction enzyme (never more than 10% final volume)
20 µl    total volume

Once the restriction digest is set up, incubation of the restriction digest is carried out at the appropriate temperature for at least 30 minutes.

Loading of the restriction digests in a gel requires addition of a loading dye. Typically, 2 ml of the 10X loading dye is added to bring the total volume up to 22ml.

DNA 10X loading dye
For 10 ml :
30% glycerol (v/v)
Trace amounts of bromophenol blue
Trace amounts of xylene cyanol FF

H. Preparation of Agarose Gel and Gel Electrophoresis

Agarose gels are usually used in electrophoresis to separate DNA fragments of different molecular weights. In order to improve the resolution of the different DNA fragments, the percentage of agarose in a gel can be altered. Unless otherwise stated, all gels used to verify restriction digests contain 0.7% agarose.

1. The gel mold was first taped at both ends. The appropriate comb that makes the wells are then placed in the slot on the gel mold.
2. 400 ml of 0.5X TBE buffer was prepared by diluting 40 ml of the 5X TBE stock solution with 360 ml of distilled water. A 50 ml aliquot was then set aside in a fresh beaker for the agarose gel.
3. To make a 0.7% agarose gel, 0.35g of high grade electrophoresis agarose was weighed out in 50 ml of 0.5X TBE buffer and then microwaved for 50 seconds to melt the agarose. Saran wrap was used to cover the flask opening.
4. The molten agarose was cooled down by running tap water over the flask.
5. 1 ml of ethidium bromide was then added into the molten agarose solution. In addition, 7 ml of ethidium bromide was also added to the remaining 350 ml 0.5X TBE buffer that was used as the electrophoresis running buffer.
6. The molten agarose was then poured into the gel mold and left to solidify for 30 minutes.
7. Once the gel solidified, the tape at both ends was pulled off and the comb carefully removed. The entire gel on the gel mold was then placed on the platform in the electrophoresis gel box. The running buffer was then poured into the box, submerging the entire gel.
8. The DNA samples to be verified were then loaded into the wells. Each DNA sample contained a 1X loading dye.
9. Since DNA is negatively charged, migration will occur from the negative terminal to the positive terminal. Thus, the electrophoresis apparatus was set up to ensure proper polarity of the terminals.
10. The gels were usually ran at 25 mA for roughly 1.5 hours. A handheld UV lamp was used to periodically check the migration of the DNA fragments in the wells.
11. Once the proper migration distance was achieved, the electrical current was discontinued and the gel was viewed under UV lamp in a transilluminator. A thermal printer was then used to capture the desired image of the gel.

I. Electroelution

Electroelution is carried out to extract the DNA from an agarose gel. Since the DNA sample has been separated based on their molecular weights, the desired DNA fragment can be isolated from the gel through electroelution. This process of segregating the targeted DNA from the rest of the DNA in the gel is known as gel purification. Though there are several ways to gel purify DNA fragment(s), electroelution using a "V" apparatus was used the lab.

1. A 500 ml elution buffer (10 mM Tris-HCl pH 8.0, 1 mM EDTA, 5 mM NaCl) was prepared and degassed for 10 minutes using a sidearm flask attached to an aspirator.
2. The electroelution apparatus was taped on to the lab bench to avoid accidentally moving the apparatus during the elution process.
3. The elution buffer was then poured into the electroelution apparatus until it covered all the wells, but was prevented from overflowing across the bridge. The gates that lead to the wells were left open.
4. Agarose gel slices which contained the desired DNA fragments were placed within the wells on the bridge.
5. A pasteur pipet was then used to flush out the "V" shaped trough with the elution buffer.
6. 100 ml of 7.5M ammonium acetate was carefully pipetted into the "V" shaped trough. Due to the density of the ammonium acetate, it sank to the junction of the "V" trough.
7. The gel slices were slid as close to the opening of the "V"  trough as possible and the apparatus was ran for 20 minutes at 150 Volts.
8. Once completed, the power supply was turned off and the gates that lead to the "V" trough were closed.
9. The buffer was then quickly and carefully removed using a syringe starting with the buffer from the side opposite of the gates. The buffer was lowered until it was below the "V" trough.
10. Excess buffer was then removed from the wells on the bridge with a pasteur pipets. The gel slices should no longer contain any DNA. A UV Lamp can be used to verify that assumption.
11. Using a special, flexible pipet tip, the ammonium acetate cushion which now contains the DNA was carefully removed and placed within a fresh 1.5 ml microfuge tube.
12. The "V" trough was rinsed once with 150 ml of elution buffer and the wash was added to the DNA / ammonium acetate solution in the 1.5 ml microfuge tube.
13. 50 ml of ammonium acetate and 1 ml of carrier glycogen (20 mg/ml) was added to the pooled DNA sample in that order.
14. 800 ml of 100% ethanol was added to precipitate the DNA. The tube was then placed in either a -70° C freezer for 30 minutes or a -20° C freezer overnight.
15. The precipitated DNA was then centrifuged for 10 minutes at 12,000 rpm. The supernatant was discarded and the DNA pellet was washed with ice cold 70% ethanol and then dried with a Savant DNA 110 Speed Vac^TM.
16. The DNA pellet was then resuspended in the appropriate amounts of TE buffer.
 

J. Quantitation of DNA

DNA absorbs UV light at a wavelength of 260 nm. Based on this knowledge, the concentration of DNA in a solution can be determined by using a UV-Vis spectrophotometer.

1. The UV-Vis spectrophotometer including the UV bulb was turned on and allowed to warm up for at least 15 minutes. The spectrophotometer was then adjusted to a wavelength of 260 nm.
2. The DNA sample was then diluted by 400 fold with the final volume being 400 ml.
3. The spectrophotometers were then blanked by inserting the quartz cuvettes which contains TE buffer into both the reference and sample cuvette holders. TE buffer was used because the DNA samples were all diluted in TE buffer.
4. Once the spectrophotometer was blanked, the diluted DNA sample can then be pipetted into the sample cuvette.
5. The optical density (OD) reading at 260 nm was taken.
6. Concentration of the DNA in the sample can then be determined by plugging the absorbance reading into the following equation:

     OD260 x 50 mg/ mL x  400 = concentration of DNA (mg/ml)

Note that the constant 50 mg/ml applies only to solutions containing double stranded DNA. For single stranded DNA or RNA, 40mg/ml should be used. For single stranded oligonucleotides, 20 mg/ml should be used.

K. QIAGEN Plasmid Midi Protocol
(Protocols from QiagenÇ GmbH and QiagenÇ Inc. Plasmid Mini Protocols)

The Qiagen midi protocol is a plasmid prep designed to isolate and purify up to 100 mg of plasmid DNA using the Qiagen-tip 100.

1.  Cell cultures of the appropriate E. coli cells containing the desired plasmid were grown overnight. 25 ml of E. coli LB culture was grown in the experiment.
2.  The bacterial pellets were centrifuged and resuspended in 4 ml of Buffer P1.
3.  4 ml of Buffer P2 was added, mixed and incubated at room temperature for 5 minutes.
4.  4 ml of chilled Buffer P3 was added, mixed and incubated on ice for 15 minutes.
5.  The mixture was then centrifuged at 13,500 rpm (20,000 g) with the Beckman JA-20 rotor for 15 minutes at 4° C. The supernatant was transferred to a new tube.
6.  The supernatant from step 5 was then centrifuged at 13,500 rpm (20,000 g) with the Beckman JA-20 rotor for another 15 minutes at 4° C. The new supernatant was transferred into a fresh tube.
7.  A Qiagen-tip 100 was then equilibrated with 4 ml of Buffer QBT.
8.  The supernatant from step 6 was applied to the equilibrated Qiagen-tip 100.
9.  The Qiagen-tip 100 was then washed with 2 x 10 ml Buffer QC.
10. The DNA from the tip was eluted with 5 ml of Buffer QF.
11. The eluted DNA was then precipitated with 0.7 volumes of isopropanol and then centrifuged at 11500 rpm (15,000 g) for 30 minutes at 4° C.
12. The DNA pellet was then washed with 2 ml of 70% ethanol and then dried with the Savant DNA 110 Speed Vac^TM.
13. The pellet was resuspended in a suitable volume of buffer.

 L. QIAGEN Plasmid Maxi Protocol
(Protocols from QIAGENÇ GmbH and QIAGENÇ Inc. Plasmid Maxi Protocols)

The Qiagen midi protocol is a plasmid prep designed to isolate and purify up to 500 mg of plasmid DNA using the Qiagen-tip 500.

1. Cell cultures of the appropriate E. coli cells containing the desired plasmid were grown overnight. 100 ml of E. coli LB culture was grown in the experiment.
2. The bacterial pellets were centrifuged and resuspended in 10 ml of Buffer P1.
3. 10 ml of Buffer P2 was added, mixed and incubated at room temperature for 5 minutes.
4. 10 ml of chilled Buffer P3 was added, mixed and incubated on ice for 20 minutes.
5. The mixture was then centrifuged at 13,500 rpm (20,000 g) for 30 minutes at 4° C. The supernatant was transferred to a new tube.
6. The supernatant from step 5 was then centrifuged at 13,500 rpm (20,000 g) with the Beckman JA-17 rotor for another 15 minutes at 4° C. The new supernatant was transferred into a fresh tube.
7. A Qiagen-tip 500 was then equilibrated with 10 ml of Buffer QBT.
8.  The supernatant from step 6 was applied to the equilibrated Qiagen-tip 500.
9. The Qiagen-tip 500 was then washed with 2 x 30 ml Buffer QC.
10. The DNA from the tip was eluted with 15 ml of Buffer QF.
11. The eluted DNA was then precipitated with 0.7 volumes of isopropanol and then centrifuged at 11500 rpm (15,000 g) for 30 minutes at 4° C.
12. The DNA pellet was then washed with 5 ml of 70% ethanol and then dried with the Savant DNA 110 Speed Vac^TM.
13. The pellet was resuspended in a suitable volume of buffer.

M. Quick Screen Gel Electrophoresis

The quick screen gel electrophoresis is carried out to screen bacterial colonies for the presence of plasmids without having to carry out plasmid mini-preps. If a transformation produced too many colonies, the quick screen will allow us to identify the colonies that might have plasmids as well as whether the plasmids contain the desired insert.

1. The bacterial colonies were picked with sterile yellow pipet tips and transferred to  numbered microfuge tubes containing 50 ml of LB (Luria Broth) + antibiotic.
2. The microfuge tubes were placed in the incubator for 2 hours at 37° C.
3. 40 ml of the cell culture was drawn from each numbered tube and placed in a fresh 500 ml microfuge tube containing 40 ml 2X Quick Dye. The tube was thoroughly mixed and the samples were ready for loading.
4. The protocols for making the agarose gels in the quick screen method is similar to that described in section H. (Preparation for agarose gel and gel electrophoresis). However, since the gel is larger, 300 ml of 0.5X TBE buffer was used to make the gel and 1 L of running buffer was also prepared.
5. Once all the samples were loaded into the wells, the gel was ran at 25 mA for almost 2 hours before viewing the it under the UV transilluminator. The desired image was then captured and printed on the thermal printer.

Note : Loading of samples into the wells are tricky since the samples are very viscous. Scraping the pipet tip at the edge of the well before lifting the tip up will prevent the sample from escaping the well and thus disperse in to the running buffer.

2X  Quick Dye
5.0 ml 1M NaOH
1.0 ml 0.5 M EDTA pH 8.0
0.5 g SDS or 2.5 ml 20% SDS
0.025 g bromophenol blue
5.0 ml glycerol
dH₂O up to 50.0 ml

Autolysin is produced by C. reinhardtii during mating of the plus and minus gametes. Its main function is to break down the cell walls during mating so that fusion of the mating types can occur. Autolysin is harvested from mating cells and used in transformation of C. reinhardtii. Since plasmids cannot penetrate cell walls, it is crucial that the cell walls are broken down prior to transformation.

1. Both R3⁺ and NO^- strains were plated on 10 (100 ml) TAP plates each and allowed to grow.
2. Each plate was then flooded with 12 ml of sterile NFHSM and put under lights for about 1.5 hours.
3. The cells were then carefully scraped with a sterile rod.
4. The R3⁺ plate was then poured on to a NO^- plate on a 1:1 ratio. The plates were then sloshed around to mix the two cells together.
5. Plates were then returned to the light shelf for another 20 minutes.
6. The plates were then poured into a sterile 250 ml centrifuge bottle and centrifuged for 10 minutes at 9500 rpm (10,000 g).
7. The supernatant was then aliquoted into 2.5 ml samples in sterile plastic tubes and stored in the -70° C freezer.
8. A 200 ml aliquot of the autolysin was plated on a TAP plate to check for the presence of any bacterial or fungal colony.
9. The autolysin was then tested to ensure that it worked. The autolysin was mixed with wild type gametes and vegetative cells for an hour under lights. An aliquot was then mixed with the  0.2% NP-40 and cell debris was observed under the light microscope.
 

NFHSM
To make 1 L:
4 ml of Ca²⁺/Mg²⁺ stock solution
8 ml of phosphate stock solution
1 ml of trace elements
up to 1 L ddH₂O
Autoclave

1. Chlamydomonas cells were first grown in liquid TAP medium for several days to achieve a density of roughly 1.5 x 10⁶ to 3 x 10⁶ cells/ml. Cells were either incubated under the light shelf at room temperature or in the lighted growth chamber at 30° C.
2. For co-transformations that involved the nit1 plasmid pMN56, the cells were transferred to an NH₄⁺ free TAP medium for 4 hours and placed on a shaker in the light shelf.
3. Using a hemocytometer, the number of cell/ml in the culture was calculated.
4.  2.5 x 10⁷ to 6 x 10⁷ cells were transferred to a sterile 50 ml blue capped tube.
5. Using a clinical centrifuge, the cells were spun down at 1435 rpm (700 g) for 7 minutes
6. The supernatant was quickly discarded and the cells were resuspended in 2 ml of sterile autolysin and placed under lights for 1 hour.
7. An aliquot of the autolysin incubated cells was then obtained and the cells were tested for cell wall loss with 0.2% NP-40.
8. The cells were then spun down in a clinical centrifuge at 1435 rpm (700 g) for 5 minutes and the supernatant was discarded.
9. To the pellet, 300 ml of NH₄⁺ free TAP medium, 1 mg of pMN56 and 100 ml of 20% PEG (filter sterilized) were added in the specific order. For co-transformations, an additional 5 mg of the desired plasmid to be transformed was added. The solution was then mixed by tapping the tube gently.
10. 300 mg of sterile 0.5 mm size 6 glass beads were also added to the tube which was then vortexed at top speed for 15 seconds.
11. 800 ml of NH₄⁺ free TAP medium was then added to the tube and mixed by gentle tapping.
12. The transformants were then plated on 3 NH₄⁺-free TAP plates in 400 ml aliquots.

Calf intestine alkaline phosphatase (CIAP) is an enzyme that catalyzes the hydrolysis of the 5í end phosphate groups of DNA and RNA molecules. As a result, recircularization of DNA is prevented due to the absence of the 5í phosphate groups.

1. CIAP is incubated with the linearized plasmid vector to remove the 5í end phosphate groups.
2. The stock CIAP was diluted with 1X reaction buffer to achieve a final concentration of 0.01 U/ml.
3. The amount of CIAP used was calculated based on the requirement that each pmol of DNA ends require 0.01 U of CIAP. (1mg of 1000 bp DNA = 1.52 pmol DNA = 3.03 pmol of DNA ends).
4. The DNA to be dephosphorylated was then purified via ethanol precipitation and resuspended in 40 ml TE buffer.
5. A CIAP reaction mixture was then prepared :
 
REAGENTS                                  VOLUME  (ml)
 DNA (up tp 10pmol of 5í-ends)              40
 CIAP 10X reaction buffer                        5
 Diluted CIAP (0.01 U/ml)               up to 5
 Total                                                    50

6.   The reaction mixture was then incubated at 37* C for 30 minutes.
7.   Another aliquot of CIAP equivalent to the same amount used in the mixture was added and the reaction mixture was incubated at 37° C for another 30 minutes.
8.   300 ml of CIAP stop buffer was added after the incubation period. The DNA was then
cleaned via phenol/chloroform/isoamyl alcohol extraction and then precipitated with 100% ethanol (with the addition of 0.5 volumes of 7.5M ammonium acetate at pH 5.5).

CIAP Stop Buffer
10 mM  Tris-HCl, pH 7.5
1 mM  EDTA, pH 7.5
200 mM  NaCl
0.5%  SDS

 Q. Transformation with Epicurian Coli XL10-Gold^TM Ultracompetent Cells
(Protocols modified from StratageneÇ Epicurian Coli XL10-Gold^TM Ultracompetent Cells instruction manual)

1. The XL10-Gold ultracompetent cells were first thawed on ice.
2. The cells were then gently mixed by hand and a 100 ml aliquot of cells were placed into a prechilled 15 ml Fisher brand polypropylene tube.
3. 4 ml of the beta-mercaptoethanol provided in the kit was then added into the aliquot of cells
4. The tube was then swirled gently and the cells were incubated on ice for 10 minutes with gentle swirling every 2 minutes.
5. 5 ml of the ligation reaction was then added into the tube.
6. The tubes were then incubated on ice for 30 minutes.
7. The tube was then heat-pulsed in a 42° C water bath for 30 seconds.
8. The tube was then incubated on ice for an additional 2 minutes.
9. 0.9 ml of the preheated NZY⁺ broth (at 42° C) was added to the tube and the tube was incubated at 37° C for 1 hour with shaking at 225-250 rpm.
10. The cells were then spun down at 12,000 rpm for 20 seconds and the supernatant was discarded. The pellet was then resuspended with 20 ml of NZY⁺ broth and then plated on an appropriate antibiotic plate.
 

NZY⁺ Broth (per Liter)
10.0 g of NZ amine (casein hydrolysate)
5.0 g of yeast extract
5.0 g of NaCl
Adjusted the pH to 7.5 with NaOH
Autoclaved
The following supplements were added prior to use :
12.5 ml of 1 M MgCl₂ and 12.5 ml of 1M MgSO₄
20.0 ml of 20% (w/v) filter-sterilized glucose solution