Protocols

Making cells competent

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Most of the time, we used Top10 chemically competent cells. We did make a stock of chemically competent DB3.1 cells with the following protocol (found on OpenWetWare). We found that these cells were indeed very competent.

You will need TSS buffer, for 50 mL:

• • 5g PEG 8000
  • 1.5 mL 1M MgCl2 (or 0.30g MgCl2*6H20)
  • 2.5 mL DMSO
  • Add LB to 50 mL

Filter sterilize (0.22 μm filter) TSS buffer and store at 4ºC or -20ºC

Preparing the cells:

• Grow a 5ml overnight culture of cells in LB media.
• In the morning, dilute this culture back into 25-50ml of fresh LB media in a 200ml conical flask. You should aim to dilute the overnight culture by at least 1/100.
• Grow the diluted culture to an OD600 of 0.2 - 0.5. (You will get a very small pellet if you grow 25ml to OD600 0.2)
• Put eppendorf tubes on ice now so that they are cold when cells are aliquoted into them later. If your culture is X ml, you will need X tubes. At this point you should also make sure that your TSS is being chilled (it should be stored at 4oC but if you have just made it fresh then put it in an ice bath).
• Split the culture into two 50ml falcon tubes and incubate on ice for 10 min.

All subsequent steps should be carried out at 4oC and the cells should be kept on ice wherever possible

• Centrifuge for 10 minutes at 3000 rpm and 4oC.
• Remove supernatant. The cell pellets should be sufficiently solid that you can just pour off the supernatant if you are careful. Pipette out any remaining media.
• Resuspend in chilled TSS buffer. The volume of TSS to use is 10% of the culture volume that you spun down. You may need to vortex gently to fully resuspend the culture, keep an eye out for small cell aggregates even after the pellet is completely off the wall.
• Add 100 μl aliquots to your chilled eppendorfs and store at − 80oC.

Transformations

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Standard transformation procedure

• Remove competent cells from -80, let thaw for 10 min on ice and aliquot in 50 ul amounts.
• add 2-5 ul of vector, usually in H2O, to 50 ul cells, no mixing by pipet due to shear induction.
• keep on ice for 20 minutes (vector spreading through volume)
• heat shock (42°C) for 45 seconds
• keep on ice for 2 minutes
• add 200 ul SOC, put on 37°C for 1 hour or longer with agitation.
• plate out 250 ul on appropriate antibiotics.

Prepering chemically competant cells - TMF Buffer

Materials

• Plate of cells to be made competent
• TMF buffer
• LB media
• Ice

Glassware & Equipment

• Falcon tubes
• 500μl Eppendorf tubes, on ice
• 200ml conical flask
• 200μl pipetman or repeating pipettor
• 5ml pipette

Preparation

1. Grow a 5ml overnight culture of cells in LB media. In the morning, dilute this culture back into 40ml of fresh LB media with 0.8 ml of Mg-mix (0.5M Magnesium chloride + 0.5M Magnesium sulfate) in a 100ml conical flask. You should aim to dilute the overnight culture by at least 1/100.
2. Grow the diluted culture to an OD₆₀₀ of 0.5 - 0.8.
3. Put eppendorf tubes on ice now so that they are cold when cells are aliquoted into them later. If your culture is X ml, you will need X tubes. At this point you should also make sure that your TMF is being chilled (it should be stored at 4°C but if you have just made it fresh then put it in an ice bath).
4. Split the culture into two 50ml falcon tubes and incubate on ice for 10 min.

All subsequent steps should be carried out at 4°C and the cells should be kept on ice wherever possible

1. Centrifuge for 15 minutes at 4000 rpm and 4°C.
2. Remove supernatant. The cell pellets should be sufficiently solid that you can just pour off the supernatant if you are careful. Pipette out any remaining media.
3. Resuspend in 4ml chilled TMF buffer and add 1 ml of 40% glycerol. You may need to vortex gently to fully resuspend the culture, keep an eye out for small cell aggregates even after the pellet is completely off the wall.
4. Add 100 μl aliquots to your chilled eppendorfs.
5. Flash freeze the eppendorfs containing the cells with liquid nitrogen.
6. Store the cells at -80°C.
7. It is a good idea to run a positive control on the cells.

Restrictions

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Try to do a restriction in a relatively large volume. As a rule of thumb, use a volume of 50 ul / 500 ng DNA.

• Calculate the amount of DNA you want to use
• add H2O
• add 10 x H buffer (Roche)
• add your calculated amount of DNA
• add 0.5 ul of each enzyme. Keep in mind 0.5 ul = 5 U, where 1 U is defined as the amount of enzyme cutting 1000 ng of DNA / hour, so for extremely large amounts of DNA adjust this.
• keep on 37°C for 2-3 hours.

Purifying small DNA parts

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Protocol found on OpenWetWare

This protocol is for a simple ethanol precipitation of small fragments. This protocol was used to (partially) purify a DNA fragment containing a ribosome binding site (~40 bp) during 3A assembly]. The fragment was generated via restriction digest and it was used in a ligation reaction. Note that this protocol simply concentrates your sample and removes enough salts/enzymes for ligation to be successful. All DNA fragments from your digest will still be present in your pellet. These residual DNA fragments do not matter for 3A assembly which selects against incorrect ligation products.

Materials

• Absolute Ethanol (100% = 200 proof)
• 95% ethanol
• Tabletop centrifuge
• -80°C freezer

Procedure

1. Add 2 volumes ice cold absolute ethanol to sample.
   Generally the sample is in a 1.5 mL eppendorf tube. I recommend storing the absolute ethanol at -20°C.
2. Incubate 1 hr at -80°C.
   The long incubation time is critical for small fragments.
3. Centrifuge for 30 minutes at 0°C at maximum speed (generally >10000 g at least).
4. Remove supernatant.
5. Wash with 750-1000 μL room-temperature 95% ethanol.
   Another critical step for small fragments under 200 base pairs. Generally washing involves adding the ethanol and inverting several times.
6. Centrifuge for 10 minutes at 4°C at maximum speed (generally >10000 g at least).
7. Let air dry on benchtop.
   I generally let the pellet air dry completely such that it becomes white so that all residual ethanol is eliminated.
8. Resuspend in an appropriate volume of H₂O.
   Many protocols recommend resuspending in 10 mM Tris-HCl or TE. The advantage of TE is that EDTA chelates magnesium ions which makes it more difficult for residual DNases to degrade the DNA. I generally prefer H₂O and don't seem to experience problems of this sort. If you plan to ultimately use electroporation to transform your DNA then resuspending in H₂O has the advantage of keeping the salt content of your ligation reaction down.

DNA precipitation

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Standard Qiagen Minprep Kit was used for plasmid isolation.

Ligation

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First make sure you have purified the DNA after restriction. Ligation should be in a small volume (we usually use 15 ul), so elute your DNA from the column in a small volume/high concentration.

• add H2O
• add 10 x ligation buffer
• add backbone and insert (theoretically in a 1:3 or 1:4 ratio, for 3A assembly it seemed to work at 1:1 ratios, possibly even better). DNA amounts added are at least 50 ng of the backbone and if possible 100-150 ng of the insert DNA (including it's backbone).
• add 1 ul of T4 Ligase.
• keep the reaction at 16ºC for at least 2 hours, but o/n is preferable.
• if used for transformation, all DNA can be added to competent cells, or if you want to analyze it on gel, keep 5 ul.

PCR

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Colony PCR

• Make biobrick mastermix, containing per sample:
  • 12.5 ul Taq mastermix
  • 2.5 ul 10x forward biobrick primer
  • 2.5 ul 10x reverse biobrick primer
  • 7.5 ul H2O
• Put 25 ul in the PCR tubes.
• With a toothpick or pipet point, touch a colony and stir it through the fluid
• Run the iGEM colpcr program (to be added later)

PCR using Taq Mastermix

Contents of the PCR mix is the for a large part the same as mentioned above for the Colony PCR. Differences will be noted here. First, instead of biobrick primer, any primer of choice can be added, also 2.5ul if standard solution has a concentration of 10 pmol/ul. Also x ul template DNA from a sample is added, where x depends on the total concentration of DNA in the sample. Typically 50 to 100 ng of total DNA is added. 7.5 - x ul of H₂O is added to the mix.

PCR program is:
1. 5' @ 95ºC
2. 1' @ 95ºC
3. 1' @ annealing temperature of the primer
4. 1' @ 72ºC (1' is long enough for 1kb, longer times can be used if larger products are formed)
5. repeat steps 2-4 29x (total of 30 cycles, more can be added if necessary)
6. 5' @ 72ºC
7. ∞ @ 4ºC (PCR can be stopped and stored in the fridge at any time from this point on)

PCR using Pfx polymerase

Mastermix does not exist for the Pfx polymerase. This means the components have to be added seperately. The mix consists of:

• x ul template DNA (again 50 - 100 ng total)
• 5.0 ul 10x buffer
• 2.5 ul forward primer (10 pmol/ul)
• 2.5 ul reverse primer (10 pmol/ul)
• 0.2 ul Pfx
• 1.5 ul dNTP's (10 mM)
• 1.0 ul MgSO₄ (50 mM)
• 37.3-x ul H₂O

The PCR program looks the same as mentioned above for Taq polymerase, only difference is the elongation temperature in step 4. This is 68ºC for Pfx.

Gradient PCR

Gradient PCR is mainly used to determine the best annealing temperature for primers. This is done in this project with Taq polymerase mastermix, as this is cheaper than Pfx. However, as long as a PCR machine capable of making gradients is present, a gradient PCR can be performed with any polymerase. During the annealing step (step 3 in the taq mastermix protocol) every column in the PCR machine has a different temperature, going up from left to right. The range of the gradient can be installed manually, however the actual temperatures cannot (at least not in our machine). An example of PCR products put on gel after a gradient PCR can be seen in the lab notebook at the 20th of August, where gradients of 5ºC in 12 steps were tested for the atoB, idi and ispA primer pairs.

DNA gels

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• Take a flask of 0.8% up to 1.5% molten agarose from the 70oC stove.
• Pour a it in a taped gel tray.
• Add ca. 5 ul of SYBRSafe (depending on size gel)
• Add a comb and let the gel harden for ca. 15 minutes.
• Remove the comb and the tape and put the gel tray in an electrophoresis tray.
• Add enough 1x TBE to completely cover the gel.
• Add DNA loading buffer to your samples and load them.
• Let the gel run at a voltage between 60V and 120V, depending on desired resolution/time available.
• Visualize the DNA by putting it in the imager for taking a picture, or if you want to cut out your DNA, put it on the blue light emitter.

Fluorescence Measurements

Bacterial transformation

Introduction

Transformation is the process of introducing foreign DNA (e.g plasmids, BAC) into a bacterium. Bacterial cells into which foreign DNA can be transformed are called competent. Some bacteria are naturally competent (e.g B. subtilis), whereas others such as E. coli are not naturally competent. Non-competent cells can be made competent and then transformed via one of two main approaches; chemical transformation and electroporation.

There are advantages and disadvantages to both transformation methods. In general, chemical transformation is less prone to error and faster however electroporation produces a higher transformation efficiency (fraction of transformed cells that actually uptake the foreign DNA). See Molecular Cloning for a fuller discussion of both approaches.

Protocols

OpenWetWare already has a number of protocols relating to bacterial transformation but more are always welcome.

If you use a variant on one of these protocols please feel free to add a link to your protocol from one of these pages so other users can find a protocol that works for them. Additionally, if anyone uses the Innoe or Hanahan high-efficiency protocols, then please add protocols here.

Chemical transformation

If you plan on doing a chemical transformation, then you should see these pages -

• Preparing chemically competent cells
• Preparing TSS buffer
• Transforming chemically competent cells
• Preparing chemically competent cells (Inoue)
• Transforming chemically competent cells (Inoue)
• TOP10 chemically competent cells

Chemical transformation buffer comparison

Someone should check out the claims of Nishimura90. tk 08:58, 25 September 2007 (EDT)

Rubidium chloride transformation protocol [http://wheat.pw.usda.gov/~lazo/methods/goldberg/competen.html here]

Someone should check the claims of 1e10 chemical competence using 10% ethanol and calcium chloride protocols [http://www.ejbiotechnology.info/content/vol10/issue1/full/10/index.html here].

Electroporation

If you plan on using electroporation, then see these pages -

• Electrocompetent cells
• Electroporation

References

Medium preparation

Buffers & (Stock) Solutions

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Antibiotics (1000x stock solutions)

• Ampicillin: 100 mg/ml in H₂O
• Chloroamphenicol: 34 mg/ml in etOH
• Kanamycin: 10 mg/ml in H₂O
• Tetracycline: 5 mg/ml etOH

SOB (Super Optimal Broth)

For 1 liter dissolve in H₂O

• 20 g Bacto tryptone
• 5 g Bacto-Yeast extract
• 0.5 g NaCl
• 10 ml 250 mM KCl
• adjust pH to 7.0
• before use add 5 ml of 2mM MgCl₂

SOC (Super Optimal broth with Catabolite repression)

• add 20 mM glucose to 1L SOB.
• You can also order small bottles from Invitrogen (which is what we did)

LB medium (Lysogeny Broth^{[http://jb.asm.org/cgi/content/full/186/3/595]}, but better known as Luria-Bertani Medium)

In 950 mL H₂O

• 10 g Bacto Tryptone
• 5 g Bacto-Yeast extract
• 10 g NaCl
• adjust pH to 7.0

10x TBE (Tris, Boric Acid, EDTA)

To make 1L, dissolve in 950 ml H₂O

• 54 g Tris
• 27.5 g Boric Acid
• 4.65 g EDTA or 20 ml 0.5M EDTA pH 8.0

6x DNA Gel loading buffer

• Dissolve in H₂O
• 0.25% Bromophenolblue
• 0.25% Xylene Cyanol FF
• 40% (w/v) Sucrose

10x PBS (Phosphate Buffered Saline)

In 950 mL H₂O dissolve:

• 11.5g Na₂HPO₄
• 2g KH₂PO₄
• 80g NaCl
• 2g KCl

Adjust volume to 1L
The pH of 1x PBS should be 7.4

Source

Adapted From:

J. Sambrook, D.W. Russell, Molecular Cloning: A Laboratory Manual (Cold Spring Harbor Laboratory Press, New York, ed. 3, 2001) pg. A2.2

Lock/key synthesis

Protocol for Lock/key synthesis

Conjugation Protocol

See Conjugation Protocol.