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Protocols

Contents

1. Making cells competent
2. Transformations
3. Restrictions
4. Purifying small DNA parts
5. DNA precipitation
6. Ligation
7. PCR protocols
8. DNA gels
9. Luciferase assays
10. Protein precipitation
11. Cell lysis
12. Buffers & (stock) solutions

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.

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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Another protocol for DNA precipitation, it was used to concentrate DNA samples for sequencing.

• Add 1/10 volume of 3M Sodium Acetate (NaAc), pH 4.8
• Add 2 volumes of 96% ethanol (EtOH)
• Store for at least 1h @ -20ºC or 20' @ -80ºC (can also be stored o/n)
• Spin for 20' at max speed and 4ºC
• Decant supernatant and wash pellet with 1.5 volume of 70% EtOH (EtOH has to be cold)
• Spin for 10' at max speed and 4ºC
• Decant supernatant and air-dry pellet in approximately 15' (no EtOH should be left)
• Resuspend pellet in wanted volume of H₂O or TE
• Incubate for 10' @ 4ºC to ensure all DNA is dissolved
• NanoDrop for concentration and store at -20ºC for later use

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.

Touchdown PCR

Some of the ordered primers had long sequences that are not supposed to bind to the target DNA (the pre- and suffix for forward and reverse primer, respectively). Here low annealing temperatures could lead to a lot of aspecific product formation, while high annealing temperatures could be too specific, causing very little product formation. To suppress this, a touchdown PCR can be performed. Again 50 - 100 ng of template DNA should be used and any polymerase. The PCR program used in this project, with Pfx polymerase, looked like this:
1. 5' @ 94°C
2. 1' @ 94°C
3. 1' @ 65°C --> temperature is lowered with 0.5°C per cycle
4. 3' @ 72°C
5. go to 2, 20 cycles in total
6. 1' @ 94°C
7. 1' @ 94°C
8. 3' @ 72°C
9. go to 6, 20 cycles in total
10. 7' @ 72°C
11. ∞ @ 10°C

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.

Luciferase Assays

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Due to some protocols not working as desired, we've used various different ones. The one listed here is the specific measurement protocol, other more detailed protocols can be found under the following links: 25th of September protocol; 15th of October protocol

Measurements

The luminometer used is the BioTek Gen5 plate reader.
The luciferase assay kit used is the Promega Renilla Luciferase Assay System.

• Mix 1 ul of 100X luciferase substrate in 100 ul of assay buffer per sample in the luminometer's reagent bottle.
• At least in duplo, put 20 ul of soluble fraction of cells in a well for all samples of a white opaque 96 wells plate.
• Put the prime plate in the plate reader(usually on top of the plate reader and surprisingly labeled 'priming plate')
• Put the bottle with assay buffer under reagent needle no. 1, making sure the tip of the needle is in a position to reach all the assay buffer (the lowest point in the bottle). Fix it in this position by the elastic rubber band.
• Rinse the tubing by priming 5 ml of H₂O on the priming plate.
• Purge the tubing for 1.5 ml, leaving empty tubing.
• Prime the luminometer with 1000 ul assay buffer in the priming plate, which should exactly fill the tubing. If not sure, you can prime 15 ul until you see a small spot of fluid on the priming plate.
• Measure luciferase activity by:
  • Adding 100 ul of assay buffer to a well in the slowest possible fill rate (225 ul/s)
  • Delay 2 seconds
  • Measure/integrate luminescence for 10 seconds.
  • Repeat for every well
  • There is a standard protocol on the computer in which you only have to indicate the wells to be assayed.
• When finished, purge the tubing (If there's any assay buffer left, it can be stored and frozen at -80ºC for short periods (1 week at most) according to the technical manual)
• Rinse the tubing with 5000 ul of ethanol, and purge it for 1.5 ml.
• Rinse the tubing with 5000 ul of H2O, and purge it for 1.5 ml.
• The tubing and injector should be clean and empty now.
• Clean your plate and mark the wells you've used/throw away the plate.

Protein Precipitation

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During the project, several ways of protein precipitation were used. Here is an overview of all of them.

Perchloric Acid (PCA)

• Add 1 volume of 1M PCA to sample and mix
• Spin for 20' @ 1,500g and 4ºC
• Remove supernatant and spin again for 20' @ 1,500g and 4ºC
• Remove the supernatant as much as possible and resuspend in wanted volume of H₂O

Acetone/Trichloric Acid (TCA)

• Mix 10 volumes of cold 10% TCA in acetone (stored @ -20ºC) with your samples, vortex, and incubate at -20ºC for at least 3h, but o/n is optimal
• Spin samples 10' @ 15,000g and remove supernatant
• Wash pellet with 10 volumes of acetone, vortex, and incubate for at least 10' at -20ºC
• Spin 5' @ 15,000g, remove supernatant (carefully) and air dry pellets
• Resuspend in wanted volume of H₂O

TCA/Deoxycholate (DOC)

• Add 1/100 volume of 2% DOC, mix, and incubate on ice for 30'
• Add 100% TCA so that final concentration of TCA in the sample is 15%
• Vortex immediately to avoid formation of large conglomerates that can trap contaminants
• Keep the sample on ice for at least 1h to allow protein to precipitate, but prefarably o/n
• Spin 10' @ 15,000g and remove supernatant as much as possible
• Wash pellet with EtOH or Acetone (stored @ -20ºC)
• Vortex and incubate at RT for 5'
• Spin for 10' @ 15,000g and remove supernatant
• Repeat the last three steps (wash pellet twice)
• Dry pellet (we let it air dry, although the original protocol suggested to do it under a SLOW stream of nitrogen)
• Resuspend in wanted volume of H₂O

Methanol (MeOH)/Chloroform

• Add 4 volumes of MeOH and vortex well
• Add 1 volume of chloroform and vortex
• Add 3 volumes of dH₂O and vortex
• Spin 2' @ 15,000g - the sample will divide in two phases, proteins should be at the liquid interface
• Remove aqueous top layer, add 4 volumes of methanol and vortex
• Spin 2' @ 15,000g
• Remove supernatant as much as possible
• Air dry pellet (again, original protocol mentioned drying under nitrogen or speed-vacuum)
• Resuspend in wanted volume of H₂O

Cell Lysis

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Promega lysis buffer

• Spin off 1ml of culture for 5' @ 10,000 rpm and 4ºC
• Decant sample and get out as much of the LB medium as possible
• Resuspend pellet in 1ml of 1x lysis buffer in H₂O
• Incubate for 30' on ice
• Spin off 2' @ max speed and 4ºC
• Transfer supernatant (with protein) to a fresh eppendorf tube

Bead beater

• Spin off 1ml of culture for 5' @ 10,000 rpm and 4ºC
• Decant sample and get out as much of the LB medium as possible
• Resuspend pellet in 1ml of 1x PBS
• Add 0.5g of small acid-washed glass beads
• Add 20ul of 2uM lysozyme
• Put samples in the bead beater for 1h in the cold room
• Spin off 2' @ max speed and 4ºC
• Transfer 600ul of supernatant (with protein) to a fresh eppendorf tube

Fastprep

• Spin off 1ml of culture for 5' @ 10,000 rpm and 4ºC
• Decant sample and get out as much of the LB medium as possible
• Resuspend pellet in 1ml of 1x PBS
• Add autoclaved glass bead (d=1mm) to the sample, the amount needed equals the amount filling the conical part at the bottom of a 2 ml Greiner Bio1 microcentrifuge tube
• Shake the sample 5s at intensity 5 in the Thermo Savant FastPrep FP120 Homogenizer
• Spin off 2' @ max speed and 4ºC
• Transfer 500 ul supernatant (with protein) to a fresh eppendorf tube

Sonication

• Spin off 1ml of culture for 5' @ 10,000 rpm and 4ºC
• Decant sample and get out as much of the LB medium as possible
• Resuspend pellet in 1ml of 1x PBS
• Sonicate samples 2 times for 15 seconds with a 15 second pause in between. Make sure samples are kept on ice during sonication.
• Spin off 2' @ max speed and 4ºC
• Transfer supernatant (with protein) to a fresh eppendorf tube

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

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