Classic Cut & Paste Cloning
Introduction
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Cloning Sensei's Guide For the Aspiring Cloning Ninja
by Karmella Haynes, 2012
updated by Cassandra Barrett, 2016
When I was a postdoc in Pam Silver's lab at Harvard (2008 - 2011), my lab mates and I generated large numbers of BioBrick assemblies so rapidly, and perhaps stealthily, that one of our colleagues in the department referred to us as "cloning ninjas." This guide is based on the MIT Registry of Standard Biological Parts suggested approach, which I've modified to make ligation-based assembly as quick and painless as possible. Let's begin.
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Materials
Reagents and Disposables
LB liquid growth medium (IMPORTANT: supplemented with appropriate antibiotic)
LB agar plates (IMPORTANT: supplemented with appropriate antibiotic)
Chemically competent cells (e.g. DH5α-Turbo, BL21, NEB10, etc.) (DH5α-Turbo is the only strain that will allow you to reduce the liquid culturing time (6 hours instead of 18 hours))
Restriction enzymes (For example: Fermentas Fast Digest enzymes)
Restriction digest buffer (For example: 10X FastDigest buffer)
Green loading dye
Agrose
1X TAE
SYBR-safe or EtBr
1kb Ladder
T4 Ligase (New England Biolabs)
Ligation buffer (2x Rapid Ligation buffer, Roche)
dH2O
SOC media
Micropipette tips
1.5 mL tubes
15 mL culture tubes
Kits
Equipment
Procedure
Warm an agar plate at 37°C for at least 20 min.
Label the plate with the bacterial strain name (e.g., DH5α), the antibiotic, the BioBrick part(s) name, your initials, and the date.
Locate the desired -80°C glycerol stock. Use a sterile wooden toothpick or plastic micropipette tip to scrape up a tiny bit of the frozen bacteria and streak the plate.
Incubate the plate at 37°C for 6 hours to grow the bacteria.
Note: this may require overnight incubation if you are using a slow-growing strain of E. coli.
Label 15 ml sterile culture tube(s) appropriately. Fill each tube with 2 ml of LB growth medium + appropriate antibiotic (e.g., 100 μg/ml ampicillin).
Using a sterile pipette tip, touch the bacterial streak (or pick up a single colony) and put the tip into the LB medium (bacterial end down).
Grow the cultures overnight in a shaking 37°C incubator.
To extract the plasmid DNA from the bacteria, perform a mini prep (refer to the Qiagen miniprep protocol). 2 ml of culture usually gives a yield of about 200 ng/μl (elution vol. = 75 μl).
In your notebook, make a table of all Assemblies. List the Constructs, Inserts (and length in bp) and Vectors (and length in bp). Include the vector-only controls that you will set up ligations for. Include information about the cut sites that will be used to assemble the fragments together and the lengths of the fragments. Finally, confirm and enter the resistance marker that is on the vector backbone (Amp = ampicillin in this example). You can find the resistance marker in the plasmid map for the vector you are using.
EXAMPLE List: In line 1, the short name for the construct is KAH100/V0120 (KAH100 in vector V0120). The Insert is a "front" BioBrick insert: KAH051 cut with EcoRI and SpeI, and it is 900 bp long. The vector is KAH050-V0120, cut with EcoRI and XbaI, and it is 3700 bp long, including KAH050 (500 bp) and the V0120 backbone (3200). The last line is the negative control. There is only one negative control here because only one vector is being used for all of the ligations in this example.
Ligation table: make a table to calculate the mass of DNA you will need to perform the Assemblies.
The table below is set up to take data from Table 1 and calculate ng DNA based on the Mass ratio (e.g. 2:1 insert:vector) and Ng DNA: Vector (e.g. 50 ng) you specify. A value of 2.0 (ratio of 2 insert to 1 vector) is recommended as a starting point. The formula used in column F is:
X ng insert = (bp insert / bp vector) x 2 x 50 ng vector
Note: 50 ng of vector is recommended as a starting point. The mass of the backbone dictates the maximum number of ligated plasmids.
In your Notebook, make a list to describe the specifications of the digests and fragment pruification. Example:
Enzymes: Fermentas FD enzymes/ buffer
Incubation: @37C/ 10 min.
Purification: Sigma (elute w/ 30 uL elution sln.)
Set up your digest reaction(s) as shown in the table below:
Mix the reactions thoroughly by flickign the tubes. Incubate the digestion reactions at 37°C for 10 minutes.
Pour an agarose gel for DNA fragment purification
Make a 0.8% gel: add 0.48 g agarose to ~60 ml 1x TAE buffer in a glass flask.
Mix by swirling and microwave for 40 seconds. Mix by swirling again (to eliminate air pockets and prevent boiling-over) and microwave for 40 seconds.
Set up a gel mold and comb. Make sure the teeth are the right size to hold 30 μL of sample.
Add 6 μl of SYBR safe stain. Mix by swirling (avoid making bubbles). Pour the gel into the gel mold. Allow it to cool until it becomes opaque.
Fill a gel electrophoresis chamber with 1x TAE.
Gently remove the comb from the gel and carefully submerge the gel into the filled electrophoresis chamber.
Carefully pipette 3-6 μL pre-made 1 kb ladder mix into the first empty well and the DNA samples into the other empty wells.
Connect the electrical leads so that the positive end is at the bottom (DNA migrates to the positive end). Run the gel at 100 V.
Stop the gel when the yellow dye (Orange G) reaches the desired place on the gel (~1 hr.).
Remove the gel from the chamber and photograph under UV light.
Use a scalpel to cut the appropriate sized band(s) from the gel, place each gel slice in a 1.5 mL tube, and purify the DNA (refer to the Qiagen gel purification protocol; elute with 30 μL EB buffer). In the gel image below, blue dashed lines border where the gel was cut to excise vector fragments (lanes 1 and 2) and insert fragments (lanes 3 and 4).
Measure the concentration of the purified fragment samples with a Nanodrop Spectrophotometer. Record the absorbance (A260), purity (A260/A280), and concentration (ng/μl) for each sample.
In your Notebook, make a list to describe the specifications of the digests and fragment pruification. Example:
1. KAH051(E/S) + KAH050-V0120 (E/S)
2. KAH052(E/S) + KAH050-V0120 (E/S)
3. KAH050-V0120 (E/S) neg ctrl
-- Incubate @ room temp/ 10 min.
Make a reaction table for the ligations.
Lgn = ligation. Volumes (ul) for individual reactions are listed under each Lgn header.
IMPORTANT: To calculate the volume of Vector, divide Ng DNA: Vector (from Table 2) by ng/ul for the vector (from Table 4).
IMPORTANT: To calculate the volume of Insert, divide ng DNA: Insert (from Table 2) by ng/ul for the vector (from Table 4).
IMPORTANT: Adjust the Total volume to be as small (without decimals) as possible so that dH2O is not a negative value. A good volume is 10 - 20 ul. Adjust higher than 20 only if necessary.
dH2O and 2x buffer volumes are calculated automatically based on the other reagent volumes and the desired Total volume.
Mix the reactions thoroughly by flicking the tubes. Incubate at room temperature for 10 minutes.
Warm selection agar plates at 37°C (one for each plasmid, plus one for a zero plasmid control) for at least 15 min.
Incubate chemically competent cells on ice just until thawed. You will need 30 μL cells per ligation.
Add 30 μL thawed cells to to 1 sterile 2.0 mL tube (per ligation).
Add the total ligation reaction to the cells in each 2.0 mL tube. Pipette up and down gently to mix the cells and DNA.
Incubate on ice for 10 min.
Heat shock: Transfer the tubes to 42°C for exactly 45 seconds (heat shock) on a heat block or water bath,
...then immediately place the tubes on ice for 1 minute.
Add 750 μL sterile SOC medium to each sample.
Close the caps tightly. Place the tubes in the shaking incubator, secured in a sideways position with tape. Incubate the tubes, with shaking, at 37°C for 45 minutes.
Pre-warm the agar plates: Incuabte the selection agar plates (one per sample) at 37°C during the 45 minute recovery period.
Pellet the cells by centrifugation at top speed for 3 minutes.
Discard the SOC supernatant into the appropriate biohazard waste container.
Resuspend the pellet in 100 μL LB medium (plus the appropriate antibiotic).
Pipette the total volume of cells onto the agar; spread using sterile glass beads.
Incubate the inverted plate(s) overnight at 37°C to get colonies.
Note: To store the colonies long term, seal the plate with parafilm and keep the plate at 4°C (inverted).
Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of vector self-ligation or selection failure.
Compare the plates to estimate the ratio of “ligation” colonies to “negative control” colonies.
If the ratio is 10:1 or greater, great job! Pick 2 colonies for separate liquid cultures (see Day 1, Grow liquid cultures). Grow for 5 - 6 hours. If the ratio is less than 10:1, pick more colonies or trouble shoot and repeat the ligation & transformation.
Miniprep the DNA from the liquid cultures (see Day 2, Extract the plasmid DNA: Qiagen Miniprep Kit)
Digest 2 uL of each DNA sample with EcoRI/ PstI and check via gel electrophoresis (1% agarose) to confirm the assembled construct size. You should see one fragment that is the backbone, and another fragment that equals the total size of the two BioBrick parts you assembled.
Level 1, Newbie: Undergraduates and unseasoned scientists can expect to spend a week to two weeks on one assembly step. You will inevitably spill something, forget a step, plan an assembly incorrectly, or mess up some other inventive way. Or you have classes and can't spend every day in the lab.
Level 2, Graduate Student: Typically have experience pipetting and handling samples well and can expect to spend 3 days per assembly. If you have no life and are super-ambitious, you can crank out an assembly cycle in two days (when Day 2 procedures are started immediately after the Day 3 procedures in a single day), and complete three assemblies in one week.
Level 3, Postdoc "Cloning Ninja": If you have no life, are super-impatient, and are trying to publish papers, you can crank out an assembly cycle in two days, and complete three assemblies in one week.
