Cloning can be the stuff of nightmares. Having standard practices in designing constructs, setting up reactions, and screening really helps a lot. Here is a bit of stuff I tend to follow.

 

A. Primer design and PCR 

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-The proper design of PCR primers is essential to avoid non-specific binding of your primers to other sites, thereby creating multiple false products.

-Use a proper tool to design and maintain your primers. Benchling and Ape are really good. Benchling also give you the ability to store them on the cloud with design of constructs.

-Make sure that you have been trained to design primers correctly. 

Few tips :

-It’s easier to use a ballpark range for all your primers. 60 to 65 is useful as gives you a good balance of being high enough and avoiding nonspecificity. But this is case-dependent.

-18-24 length is usually good.

-Check for self and cross complementarity between primers.

-Also check if there are any secondary structures in the ss Oligo.

- For more complicated PCRs like fusion, overlap extension etc. a bit of optimization and maneuvering is always required.

 

B. Standard PCR reaction 

 

-For all PCRs we use Phusion or in-house purified Pfu polymerase.

-Phusion is fast but costly and Pfu can be produced easily in-house. Pfu is also a slow enzyme but in the grand scheme of things, this may not always matter. Its good to have both enzymes in the lab though.  

A standard 100 ul master mix reaction mix, which is best prepared on ice contains the following ingredients:

 

20 ul 5x HF buffer for Phusion or Pfu buffer
2 ul dNTPs (10mM each)
1 ul template, approximately 10 ng if a plasmid (Higher amount 200-250 ng in case of genomic DNA)
0.5 ul   forward primer (100 uM stock)
0.5 ul reverse primer (100 uM stock)
75 ul milliQ water
1 ul Phusion Turbo or Pfu (In house Pfu activity has to be tested to know the units)

 

The thermocycler run program will look like this:

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2 minutes initial denaturation at 98 degrees
20 seconds denaturation at 98 degrees
20 seconds annealing at 50-70 degrees (usually 55)
X seconds elongation at 72 degrees (20 seconds/kb usually works for Phusion and around 2 min 30 sec for Pfu )

Products of this reaction can be visualized on an agarose gel and also purified using a gel purification kit

 

C. Restriction digests for cloning 

 

-One-hour magical digests are quick but can mostly leave uncut product.

-It’s sometimes better to do four hour to overnight digests, which require less enzyme, and are usually better for vector backbones, which get more efficiently cut this way.

-If possible use standard FD enzymes. Buffer compatibility can sometimes be a tricky thing to get around to.

-Like any other non-standard digest, try and research before each reaction.

Products of your restriction digest can either be column purified, if only a single product is involved, or by gel extraction, if multiple products arise.

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D. Conventional cloning. 

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Introduction to Ligation 

 

Likely the most dreaded of all steps.

-T4 ligation buffers contain ATP that is essential to the success of the reaction. This may be seen as a white precipitate when buffers are cold. Let this mix properly before use.

-Freeze-thaw of 10x ligation buffer containing ATP will result in the hydrolysis of ATP, we will add ATP separately from a 10mM stock. Alternatively, make small aliquots of the buffer on first use.

-Ligation also depends on PEG 6000, which is used for molecular crowding to increase the efficiency of the reaction.

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The composition of commercially available quick ligation buffer (1x) is:

 

66 mM Tris-HCL
10 mM MgCl2
1 mM Dithiothreitol (Also see you have the fishy smell from the buffer. DTT is also important for the ligation.)
1mM ATP
7.5% Polethylene glycol (PEG6000)
pH 7.6 @ 25°C

 

The ligation mix 

Mix the following ingredients in either a 20ul reaction mix

 

• At least 50ng of dephosphorylated vector
• 3-fold molar excess of insert   (I also tend to have 1:4 and 1:5 when possible)
• 1x ligation buffer with ATP
• add water as needed to reach the appropriate volume of your reaction
• mix all by stirring and avoid quick pipetting
• lastly, add 0.5 or 1ul of T4 ligase and incubate the mixture for at least  30 minutes at room temperature. Longer does not hurt and overnight incubation of 16 degrees can also be very helpful in some cases.

 

Always set up a control, which only contains the cut vector but no insert. This will be extremely useful to check the efficiency of ligation later on.

A hack for too many false positives.

• A restriction digestion step after ligation can help eliminate false positives.

• This will only work for vector self-ligation when gel extraction was not done.

• Use an RD enzyme from the MCS, not present in your gene, and should have been removed after your first RD.

• This only needs to be a quick 30 min reaction followed by transformation. Product purification is not needed.

 

E. Gibson assembly

Store both reaction buffers and master mix in -20C freezer

 

5x Isothermal Reaction Mix

3 ml 1 M Tris-Hcl (pH 7.5)

300 μL 1 M MgCl2

60 μL 100 mM dGTP

60 μL 100 mM dATP

60 μL 100 mM dTTP

60 μL 100 mM dCTP

300 μL 1 M DTT

1.5 g PEG-8000

300 μL 100 mM NAD

balance ddH2O

6 ml Total

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Assembly Master Mix

320 μL 5X Isothermal Master Mix

0.64 μL 10 U/μL T5 exonuclease

20 μL 2 U/μL Phusion DNA Pol

160 μL 40 U/μL Taq DNA Ligase

860 μL ddH2O

1.2 ml Total

 

1.33x Gibson Master Mix:

• Taq ligase (40u/ul): 50 ul

• 5x isothermal buffer: 100 ul

• T5 exonuclease (1u/ul): 2 ul

• Phusion polymerase (2u/ul): 6.25 ul

• Nuclease-free water: 216.75 ul

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Original protocol from: this paper, doi:10.1038/nmeth.1318

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1. PCR up your fragment of choice and gel purify

2. Thaw a 15 μl assembly mixture aliquot and keep on ice until ready to be used.

3. Add 5 μl of DNA to be assembled to the master mixture. The DNA should be in equimolar amounts. Use 10-100 ng of each ~6 kb DNA fragment. For larger DNA segments, increasingly proportionate amounts of DNA should be added (e.g. 250 ng of each 150 kb DNA segment).

4. Incubate at 50 °C for 15 to 60 min (60 min is optimal).

5. Transform as usual

 

-Ideally you have an overlap of 40 bp, 20 to 25 also works

- Dilute chemically competent cells when transforming with electroporation.

- Electrocompetence is ~ 1000x more effective so is tempting.

-When preparing the isothermal reaction mix, add the PEG slowly to liquid.

-10 - 200 ng of total DNA be used for assemblies. But less constructs the better. As you go higher in the number of constructs, efficacy may go down. I may have assembled up to four or five fragments.

-As Taq pol is used for filling up the ends, there is a potential for mutations at the DNA boundaries. Its suggested the occurrence is 1 every 50 assemblies or so.

-I have used PCRs as is (with PCR cleanup only) and gel extracted DNA in my assemblies. PCR cleanup gives more colonies (more DNA, better quality (no agarose/QG contamination)) but also has more false positives (PCR template plasmid).

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F. Transformation 

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Chemically competent cells. 

-Thaw competent cells on ice and put agar plates containing the appropriate antibiotic in a 37 degree incubator.

-After the ligation reaction, use 10-15ul to add to 100 ul of competent cells.

-Incubate the cells and ligation mix on ice for 30 minutes.

-Heat shock for 60 seconds at 42 degrees in the water bath, and immediately transfer on ice.

-Incubate on ice for 1 minute before adding 800 ul of LB medium or 300 ul of SOC medium without antibiotics.

-Recover with shaking at 37 degrees for 45 minutes and pellet down by spinning for 1 minute at 8000g at RT.

-Resuspend in fresh 150 ul fresh LB and then plate everything on a petri dish containing the appropriate antibiotic using glass beads.

-Make sure that the liquid medium has been absorbed by the plate before removing the glass beads.

-Place plates in a 37-degree incubator and be patient.

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G. Electroporation.

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Reagents : Sterile and chilled MQ water, Sterile and chilled 10% glycerol. For Mycobacteria, use add 0.05% Tween 80 to the medium and the glycerol.

I have tested the method on E. coli and M. smegmatis for transforming plasmids and linear gene fragments.

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Making Electrocompetent cells

 

Important: All steps in this protocol should be carried out aseptically

• Inoculate: Prepare flask containing 10 ml of LB medium. Pick up a single colony of cells from plate and incubate the culture overnight at 37°C with vigorous aeration (180 pm in a rotary shaker).

• Inoculate 200 ml of prewarmed LB medium with final 1% overnight bacterial culture. Incubate the flasks at 37°C with agitation. Measure the OD-600 every twenty minutes (this step will take around 1.5-2 hrs).

• Rapidly cool culture: Once the OD-600 of the culture reaches 0.6-1.0 (Molecular Cloning recommends 0.4). Rapidly transfer the flasks to the pre-made ice-water bath for 20 minutes. Swirl the culture occasionally to ensure that cooling occurs evenly. Also prechill your centrifuge bottles in ice.

Note: After this point, do not let your cells warm up past 4°C

When harvesting cells by decanting, be very careful not to disturb the pellet. Mycobacterial cells are particularly problematic in this case as they tend to not stick very well to the tube.

 If necessary, aspirate instead of decant the supernatant. Also, if the pellet seems loose, sometimes it is helpful to re-spin the cells down.

Centrifuge 1: Transfer the cultures to ice-cold centrifuge bottles. Harvest the cells by centrifugation at 1000g (2500 rpm) for 15 minutes at 4°C. Decant the supernantant and resuspend the cell pellet in equal volume of ml of ice-cold MQ. 

Centrifuge 2 (water): Harvest the cells by centrifugation at 1000g for 20 minutes at 4°C. Decant the supernatant and resuspend the cell pellet in 250 ml ice-cold DI water.

Centrifuge 3 (water): Harvest the cells by centrifugation at 1000g for 20 minutes at 4°C. Decant the supernatant and resuspend the cell pellet in 10 ml ice-cold 10% glycerol.

Note: The idea here is to get off as much salt as possible such that it does not interfere with your electroporation.

Centrifuge 4 (glycerol): Harvest the cells by centrifugation at 1000g for 20 minutes at 4°C. Carefully decant the supernatant and try to get out as much liquid as possible.

• Gently resuspend the pellet in 1 ml 10 % glycerol (for every 100 ml culture used) with gentle swirling or mixing DO NOT vortex.

 

Suggestion from Openwetware: The desired concentration is 2.5×1011 cells per mL. This corresponds to an OD-600 (after 100x dilution) of roughly 3.75. It is difficult to reach this value, but it is still important to know the concentration of cells to calculate efficiencies.

• Test for arcing: Transfer 40 ul of the suspension to an ice-cold electroporation cuvette (0.1-0.2 cm gap, on middle shelf next to electroporator) and test whether arcing occurs when an electrical discharge is applied. Place the cuvette in the green holder attached to the machine. Go to option 4, Pre-set protocols; choose bacterial; choose the correct choice for your size cuvette, probably the first option for a .1 cm cuvette. If arcing occurs, wash the remainder of the cell suspension once more with ice-cold 10% glycerol to ensure that the conductivity of the bacterial suspension is sufficiently low (<5 mEq).

• Storage: Store cells at -80°C until they are required for use. For storage, dispense 40 ul aliquots of the cell suspension into sterile, ice-cold .5 ml microcentrifuge tubes, drop into a bath of liquid nitrogen and transfer to a -80°C freezer.

• To use frozen cells: Remove an appropriate number of aliquots of cells from the -80°C freezer. Thaw the tubes on ice.

 

Materials

• Electrocompetent cells

• Plasmid DNA (from a ligation reaction) or linear gene fragment.

• Electroporation cuvette (either 1mm or 2mm gap width)

• Electroporator

• 1.5 mL eppendorf tube

• LB-agar plate with appropriate antibiotic

• 1mL SOC at room-temperature

Procedure

1. Thaw frozen cells on ice and pre chill electroporation cuvettes, DNA samples and tubes on ice. Freshly prepared electrocompetent cells may be used immediately and are recommended for linear fragments, but this may also depend on how important is the knockout target.

2. Place LB-agar plates in 37°C incubator to warm.

3. Turn on electroporator and set voltage to either 1.25 kV (1mm cuvettes) or 2.5 kV (2mm cuvettes).

4. Add cells to the cuvette and the desired amount of DNA and mix gently with a P200. Let it sit for a minute and set your p1000 to 1ml and also get the SOC prewarmed. Place cells back on ice to ensure they remain cold.

5. Tap the cuvette on the counter gently so that cells are at the bottom and to remove any air bubbles.

6. Wipe off excess moisture from outside of cuvette.

7. Place in chamber of electroporator.

8. Slide the chamber in so that the cuvette sits snugly between electrodes.

9. Pulse the cells with a shock by pressing button on electroporator.

10. Remove cuvette from the chamber and immediately add SOC. This step should be done quickly to prevent cells from dying off.

11. Transfer the SOC-cell mixture to an eppendorf tube.

12. Transfer eppendorf tube to 37°C incubator and shake to promote aeration. Incubate for 1 hr for cloning or plasmids and 2-4 hours for genome manipulation.

13. Plate transformation onto prewarmed LB-agar plate supplemented with the appropriate antibiotic. I generally plate 200μL but the appropriate plating volume depends on the efficiency of the transformation. For genome manipulation, retain the unused volume and plate after 12 hours.

14. Incubate the plate overnight at 37°C and be patient.

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