Selecting Correctly Expressing Recombinants



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Blue White Screening

Blue / White colony screening is a strategy to quickly and easily distinguish between recombinant and non-recombinant colonies. It requires a special vector and a special strain of E. coli. It is particularly helpful in tricky cloning strategies such as blunt ended cloning or DNA library preparation. One can either use the protocol below to prepare the reagents to perform the screen, or you can purchase the ready-to-use Blue White Screening Reagent (Catalog  No. B2928).

How Blue White Screening Works

The first gene in the E. coli lac operon is lacZ, which encodes β-galactosidase (β-gal). The active form of β-gal is a tetramer and hydrolyses lactose into glucose and galactose. Deleting amino acids 11-41 of β-gal (called the lacZΔM15 mutation) means the enzyme is unable to form a tetramer and is non-functional (Langley et al. 1975). It was discovered that supplying amino acids 1-59 (the α-peptide) of β-gal in trans (separately) allowed the truncated β-gal to form tetramers and function again (Ullmann et al. 1967; Langley et al. 1975). Rescuing β-gal by supplying the α-peptide in this way was termed α-complementation. Later, Vieira and colleagues (Vieira & Messing 1982) realised that α-complementation could be used to screen E. coli colonies for the presence of inserts. They cloned the α-peptide coding region into a pUC plasmid and then introduced a multiple cloning site (MCS) into the middle of that region. When a piece of DNA is ligated into the MCS, it disrupts the α-peptide, rendering the β-gal non-functional.

5-bromo-4-chloro-indolyl-β-D-galactopyranoside (x-gal) is a colourless analogue of lactose. When β-galactosidase hydrolyses x-gal, it creates a blue product (5,5'-dibromo-4,4'-dichloro-indigo). In blue white screening, an E. coli strain is transformed with a ligation reaction and spread onto agar plates containing x-gal. A blue coloured colony indicates that the α-peptide in the plasmid is intact (no insert) whereas a white colony indicates that the α-peptide is disrupted (insert present).

What you will need for Blue White Screening

  • Competent cells of an E. coli strain with the lacZΔM15 mutation. Common blue white compatible strains include all SIG10 (Catalog No. CMC0001), SIG10 F' (Catalog No. CMC0002), and SIG10 5α Competent Cells (Catalog No. CMC0007).
  • A vector with the α-peptide coding region and MCS. Common blue white compatible vectors include: pGEM-T, pBluescript, pUC18 and pUC19
  • Your ligation reaction (i.e. your insert of choice into a blue white compatible vector, above).
  • Control plasmid (e.g. pBluescript).
  • Antibiotic for selecting for the vector.
  • X-gal 20 mg/ml. X-gal can be purchased ready dissolved or as a powder (Catalog No. B4252). It may be dissolved in DMSO (Catalog No. D8418) or DMF (Catalog No. D4551) at a concentration of 20 mg/ml. X-gal must be stored at -20°C and protected from light (by wrapping foil around the stock container).

How to Prepare Blue White Screening Plates

  1. Prepare some LB agar plates containing the appropriate antibiotic to select for your chosen plasmid.
  2. Onto each plate to be used for blue white screening, spread 100 μl of 20 mg/ml x-gal stock and 100 μl of 10 mM IPTG and allow the plates to dry with the lid slightly open before use. This can be performed either next to bunsen burner on the bench or in a laminar flow hood. Using a hood may dry out the plates if they are left for too long.

Bacterial Transformation

Transform your ligation reaction(s) into competent E. coli cells as usual. Spread the transformation reaction onto an x-gal IPTG plate (prepared as above). Incubate the plate overnight at 37°C. Once the colonies have grown, the plate may be incubated at 4°C for 1 hour. This helps the blue colour to develop making it easier to discern the negative colonies.

It is a good idea to include a control. Transform an aliquot of E. coli with an intact α-peptide-containing-plasmid (pBluescript for instance). The colonies on this control plate should all be blue. If they're not, then the x-gal may not have been spread evenly or the antibiotic may not be working properly.

Note: This screen does not give any information about the direction of an insert, just its presence or absence. If the insert is quite short and maintains the frame of the α-peptide, it is possible (but unlikely) that it will produce a functional α-peptide fusion giving blue colonies even when the insert is there (false negatives). However, these colonies will likely be a lighter blue than the true negatives. It is also possible (but again unlikely) to get a white colony with no insert (false positive). This could result from nuclease degradation of the linearised vector disrupting the α-peptide before re-ligation. Therefore, it is always a good idea check your insert by sequencing too.

Blue White Screening References

  • Langley, K.E. et al., 1975. Molecular basis of beta-galactosidase alpha-complementation. Proceedings of the National Academy of Sciences of the United States of America, 72(4), pp.1254–1257.
  • Ullmann, A., Jacob, F. & Monod, J., 1967. Characterization by in vitro complementation of a peptide corresponding to an operator-proximal segment of the beta-galactosidase structural gene of Escherichia coli. Journal of molecular biology, 24(2), pp.339–343.
  • Vieira, J. & Messing, J., 1982. The pUC plasmids, an M13mp7-derived system for insertion mutagenesis and sequencing with synthetic universal primers. Gene, 19(3), pp.259–268.

DNA Miniprep Protocol

The ability to work with DNA is dependent on the ability to isolate it from bacteria. The amount of DNA that is required for analysis determines the preparation technique that is used. For small quantities of DNA (<5µg) a miniprep is performed and normally provides plenty of DNA for analysis. If more DNA is required, a maxiprep can be performed, and if even more DNA is required a Mega or Giga prep can be used.

A miniprep is most often used for determining if a bacterial clone contains the correct piece of recombinant DNA. After picking colonies (typically 5-15), and growing each one in 3-5ml of LB media overnight, the bacteria are pelleted and then lysed. The genome of bacteria is tethered to the inner surface of the plasma membrane and as such stays with the membranous portion of the cell during the extraction. Small plasmids are not tethered in this manner and can be easily isolated from the cell debris.

Many laboratories use miniprep spin column kits like the GenElute™ Plasmid Mini Prep Kit (Catalog No. PLN70), or the GenElute Five Minute Plasmid Mini Prep Kit (Catalog No. PFM50) for screening but this protocol can be used if you choose not to use a kit. It is a fast and cheap routine procedure that is based on the use of fairly standard chemicals, a waterbath, pipettes, eppendorf tubes and a microcentrifuge. However, the DNA it produces can be somewhat lower in quality and quantity than the equivalent DNA isolated using a spin column kit. For diagnostic cloning however this should not be a problem.

DNA Miniprep Protocol Reagents

Protocol

  1. Start with fresh overnight liquid cultures (A single colony picked and inoculated into 3ml LB medium the evening before).
  2. Add approximately 1.5 ml of each culture into an eppendorf tube. Number the eppendorf tubes according to the numbers written on the culture tubes.
  3. Centrifuge the tubes for 1 minute at maximum speed and then discard the supernatant by throwing it out. Make sure only minor quantities of LB medium are left with the bacterial pellets.
  4. Re-suspend the pellets in 150 µl TES by pipetting the solution up and down with a 200µl pipette close to the pellet (systematically going from one side of the pellet to the other until nothing is left sticking to the wall)
  5. Quickly add 20µl of Lysozyme solution (10mg/ml in distilled water, usually kept as small stocks frozen at -20).
  6. Incubate for 5 minutes at room temperature.
  7. Pipette 300 µl of distilled water quickly to the suspensions, it should mix while the water is sprayed into the tube. Don't shake the tubes afterwards. This should be done as quickly as possible and the tubes are immediately placed in a heat block at 73°C.
  8. Incubate for 15 minutes.
  9. Centrifuge at maximum speed (13000 rpm) for 15 minutes. Sometimes, the pellet is too big (more pellet than supernatant), if so then centrifuge for another 15 minutes, be patient, it will pellet down eventually. When the pellet is big, usually the culture was too young (less than 12 hours old).

    Pour the supernatant into another eppendorf tube (don't forget the numbering). Add 5M NaClO4 (approximately 10% of the supernatants volume, usually 10% of 300µl. If some of your tubes have less supernatant, add some TE to those so that they look like the majority). Close the lids and mix by shaking the tubes. This can be done either in the rack using the lid to stop the tubes falling out or by shaking the tubes individually.
  10. Add 400 µl isopropanol, shake as before and centrifuge for 15 minutes at maximum speed.
  11. Discard the liquid and centrifuge once more for 2 minutes. Remove the last bit of liquid with a 200 µl pipette.
  12. Dry for 15 minutes in a 37°C room or incubator with the lids open.
  13. Add 50 µl of TE and put on a shaker for 5 minutes then store the DNA or proceed to the screening colonies protocol.

For most in vitro studies in mammalian culture more DNA is required than is isolated in a miniprep. Usually after a correct clone has been identified, the DNA can be re-introduced into bacteria and isolated using a maxiprep protocol. It is also possible to keep a small portion of each of the cultures that were used in the minipreps and then grow the culture containing the correct clone overnight. This saves time by not having to re-transform the bacteria again and pick a colony.

Screening by Restriction Digestion

A restriction digestion is performed in order to determine if the clone picked contains the insert. This digest is meant as a quality control, or to test different clone recombinants, and requires only a small amount of plasmid, to be digested for a standard time (1 hour) with an amount of enzyme that is in excess. The total reaction is intended to be loaded onto an agarose gel. There is no need to do a time course as for the preparative digest.

200ng of plasmid is usually sufficient and 1 unit of enzyme in a total reaction volume of 10 µl. Remember that larger plasmids may require you to use a bit more DNA if you are cutting out small fragments. 200ng of a 3 kb plasmid is 5 times more copies than of a 15 kb plasmid and so there is 5 times less copies of any fragments you cut out.

If the plasmid is not from a relatively clean maxiprep or from a silica resin based miniprep kit then you may want to use more enzyme. If the DNA is prepared using the miniprep protocol on this website you might want to double or triple the amount of enzyme.  Common enzymes used include EcoR I (Catalog No. R6265), BamH I (Catalog No. R0260), and Hind III (Catalog No. R1137).

Reagents for a clean plasmid with 1µg/µl

  • 0.2-0.5 µl plasmid
  • 1 µl 10x Restriction buffer
  • 0.1-0.3 µl Restriction enzyme 1
  • 0.1-0.3 µl Restriction enzyme 2 (Optional)
  • Make up to 10 µl with TE (Catalog No. T9285) or nuclease free water (Catalog No. W4502)

Reagents for phenol chloroform extracted miniprep plasmid (DNA concentration much lower and usually unknown)

  • 3 µl plasmid
  • 1 µl 10x Restriction buffer
  • 0.1-0.3 µl Restriction enzyme 1
  • 0.1-0.3 µl Restriction enzyme 2 (Optional)
  • Make up to 10 µl with TE or water

Protocol

  1. Calculate the total number of samples you have to screen and then add one (n+1).
  2. Make a master mix using the guidelines above with everything except the DNA to screened. So if you have ten samples use 11 µl 10x Restriction buffer, 1.1 µl enzyme and 95.7 µl water.
  3. Add an aliquot (minus the DNA quantity) of the mastermix to the number of tubes you have samples for. In this case 10.
  4. Add the DNA and mix with the tip to make sure it digests.
  5. Incubate for 1 hour at 37°C
  6. Add 1 µl of loading dye and load onto an agarose gel.

Note: Some enzymes react at temperatures above or below 37°C. Check before you set up the reaction.

Screening Colonies by PCR

To detect which bacteria have the correct recombinant DNA, it is possible to screen the colonies from the agar plate using a PCR method without growing them overnight. One method of doing this is simply to touch a colony with a sterile tip, then dip the tip briefly into a PCR mix in a PCR tube, then dip the same tip into some media to grow them overnight. Typically, this will only work efficiently if you have a relatively small region to amplify (<500bp) to detect your recombinant plasmid, and it relies on the 98ºC stages of the amplification to lyse the bacteria to release the DNA. To ensure that the E.coli are lysed efficiently prior to performing the PCR amplification, the protocol below can be used as a more reliable alternative.

  1. Prepare 5 mg/ml Proteinase K stock solution by dissolving 25 mg PCR grade Proteinase K (Catalog No. P2308) in 5ml of T0.1E (10 mM Tris-HCl, 0.1 mM EDTA, pH 8.0). Store in aliquots at –20 ºC.
  2. For each sample prepare 25 µl lysis solution by mixing 1 µl of 5 mg/ml Proteinase K stock and 24 µl T0.1E in an eppendorf tube
  3. Pick the colony using a sterile pipette tip.
  4. Spot some cells onto a labelled LB plate (with appropriate antibiotic).
  5. Deposit the remaining cells into a 25 µl aliquot of the lysis solution.
  6. Incubate the labeled plate overnight at 37 ºC to recover colonies.
  7. Begin cell lysis by incubating the samples at 55 ºC for 15 min.
  8. Incubate the samples at 80 ºC for 15 min.
  9. Cool the samples on ice.
  10. Mix and spin down briefly.
  11. Use 1 µl of lysate in a PCR reaction.

Maxipreps Protocol

Required Reagents

  • TE 50/1 buffer (50mM Tris-HCL (Catalog No. 93362) pH 8.0, 1mM EDTA (Catalog No. E6758) pH 8.0)
  • Lysozyme (Catalog No. L6876) at 10mg/ml in water.
  • 0.5M EDTA pH 8.0
  • Ribonuclease A (Catalog No. R4642) solution at 20 mg/ml in distilled water and stored in 50µl aliquots in -20°C.
  • 10% Triton X 100 in water
  • Equilibrated Phenol (Catalog No.  P4557) (pH 8.0 with 0.1% 8-hydroxyquinoline)
  • 5M NaClO4 (Catalog No. 410241)
  • Isopropanol (Catalog No. I9516)
  • TE (10mM Tris-HCl pH 8.0; 0.1 mM EDTA pH 8.0)
  • SS34 Sorvall 50ml tubes and rotor or similar
  • 30ml Corex tubes and HB4 rotor or similar

Potential Hazard: Phenol is very dangerous and causes skin burns immediately on contact. It is toxic on contact, by inhalation and by consumption. Read and understand the SDS before using any hazardous chemicals. 

Note: Sigma Aldrich Life Science is committed to bringing you Greener Alternative Products, which adhere to one or more of The 12 Principles of Greener Chemistry.  GenElute™ Maxi Prep Kits were  Designed for Safer Chemistry. Please consider use of one of these kits instead of performing phenol-chloroform DNA extractions.

Read the protocol before starting and make sure you have all the stock solutions ready. The lysozyme solution is usually made during the centrifugation period. The rest has to be ready before you start.

  1. Make a pre-culture of 3 ml LB media first thing in the morning by picking a single colony from a fresh agar plate (ideally streaked the night before).
  2. After 3 hours the pre-culture should be slightly turbid, inoculate 800ml LB media pre-warmed to 37°C (500-fold dilution) and grow overnight (24 hours). Shorter incubations may cause problems with excess proteins and RNA in the cell lysate.
  3. Centrifuge the culture using the available infrastructure (for example 60 minutes, 3699rpm) and remove the supernatant carefully. 250ml polypropylene conical bottomed tubes work well. The supernatant will still contain E.coli so do not tip it straight down the sink.
  4. Put the tubes containing the cell pellets on ice and resuspend pellets in 8 ml of ice cold TE 50/1 (see above). Pellets must be completely resuspended by occasional vortexing and placing back on ice, takes some time (5 minutes). Transfer the cell suspension with a 10ml pipette into clean SS34 50ml tubes that are standing on ice already. All further manipulations will be done on ice! This is a good point to take a break if required.
  5. Quickly add 2.5 ml of freshly made lysozyme solution (10mg/ml) to all tubes, close the tubes and turn them quickly upside down and back at least 10 times (preferentially all tubes at once. The suspension should become slightly viscous. Do not shake the tubes, just turn them. Shaking will sheer the genomic DNA and it will contaminate your plasmid DNA.
  6. Incubate for 5 minutes on ice.
  7. Unscrew the bottles and from now on remember which cap belongs to which tube (very important because the solution is viscous and a significant portion will stick to the cap). Quickly add 2.0 ml of 0.5M EDTA pH 8.0, close the tube and turn them again as before. Again, do not shake them vigorously! Suspension will become more viscous.
  8. Incubate another 5 minutes on ice.
  9. Mix 50µl of Ribonuclease A solution (see above) with 150µl Triton X-100 10% solution and make up to 1ml with TE 50/1. Add this mixture to the tubes. Then close the tubes and turn them as before (no shaking). Suspension will become even more viscous.
  10. Incubate 30 minutes on ice, without ever touching them again. The idea is to let the detergent do the rest of the cell lysis in a very gentle way, on ice.
  11. Centrifuge at 18,000 rpm in a Sorvall SS34 rotor or similar for 60 minutes. You should get a nice pellet, but sometimes the pellet can be big with very little supernatant. This usually happens when your bacterial culture is too young. In this case, repeat the centrifugation, it should help.
  12. Transfer the clear supernatant to a clean Falcon tube. Now it is possible to interrupt the procedure for longer periods by freezing the recovered supernatants at -20°C. It is also possible to have a lunch break and keep the supernatant on ice.
  13. Add 20 ml of equilibrated phenol (see above) and shake vigorously for 1 minute, spin down at maximum speed in the swing out rotor for 20 minutes.
  14. Slowly recover the water phase (upper) and make sure you leave the interphase in the tube. Transfer the upper phase to a new tube and add 20ml of chloroform, shake again as before and spin as before for 5 minutes.
  15. Slowly recover the water phase (this time it is easier to avoid the interphase), transfer to a 30ml Corex tube (between 10 and 15ml, the volumes need to be equalised by the addition of TE 50/1), add 1ml 5M NaClO4 (10% of the water volume), mix and add 8ml isopropanol (80% of the water volume).
  16. Seal the tube with parafilm, and turn it around a few times to mix everything. Then spin down in HB4 rotor at 10,000rpm for 15 minutes.
  17. Discard the supernatant, dry the pellet by leaving the lid open on the bench or at 37°C.
  18. Re-suspend in 500 µl of TE and transfer to a sterile eppendorf tube. Avoid over drying the pellet.
  19. Store at 4°C.
  20. You will need to check the DNA on gel to assess the quantity of RNA , it is usually necessary to carry out a second RNase treatment at this point.

Note: The plasmid DNA you have isolated is ready for transfection into insect or mammalian cells using standard transfection reagents like Escort™ IV (Catalog No. L3287), Universal Transfection Reagent (Catalog No. T0956), or the Calcium Phosphate Transfection Kit (Catalog No. CAPHOS).

DNA Precipitation Protocol

Precipitating DNA can be hit and miss. Sometimes you end up the same quantity and its more concentrated and cleaner, sometimes you end up with nothing. If you do it right then it should work fine but working with low concentrations of DNA (<20ng/μl) in large volumes can be tricky.

Materials

  • 3M Sodium Acetate buffer, pH 5.2 (Ideally at 4 °C)
  • Cold 100% Ethanol (Ideally at -20°C) (Catalog No. 57023)
  • Cold 70% Ethanol in sterile dH2O (Ideally at -20°C)
  • DNA sample
  • Microcentrifuge (Ideally a refrigerated centrifuge or one kept in the cold room). Centrifuge with brake off.

Protocol

  1. Transfer DNA to a container where it occupies less than one quarter of the total volume (For example, a 1.5ml tube should have no more than 375 µL of DNA solution).
  2. Add one tenth of the DNA volume of Sodium Acetate buffer to equalize the ion concentrations.
  3. Add 2-3 volumes of cold 100% ethanol and place the sample in a -20°C freezer for at least one hour.
  4. Centrifuge sample for 15 minutes at highest speed (12-13000 rpm) in a 4°C microcentrifuge.
  5. Remove as much of the supernatant as possible with either a 1ml pipette or a glass capillary tube. Be careful not to disturb the pellet. If you get all the liquid out, go on to step 6.  If not then re-centrifuge briefly, then remove the rest with a 200 µL pipette.
  6. Add 250 µL of cold 70% ethanol.
  7. Centrifuge for 5 minutes in a 4 °C centrifuge at maximum speed.
  8. Remove supernatant with a 200 µL pipette; evaporate remaining ethanol in a 37 °C water bath. Do not do this for too long because overdrying the pellet can make it difficult to resuspend.
  9. Re-suspend pellet in desired volume of water or TE buffer.

Tip 1: Performing the precipitation at 4°C is not strictly necessary for it to work but it can increase the efficiency, especially of low concentration samples (<20ng/ µl).

Tip 2: DNA precipitation can be used to clean a DNA sample, not just concentrate it. It will remove any contaminating salts from the solution.

RNase Treatment and RNA Removal

Reagents

Potential Hazard: Phenol is very dangerous and causes skin burns immediately on contact. It is toxic on contact, by inhalation and by consumption. Read and understand the SDS before using any hazardous chemicals.

Potential Hazard: Chloroform is toxic by inhalation, oral consumption and skin contact. Read and understand the SDS before using any hazardous chemicals.

  1. Dilute the maxiprep to 5ml with TE in a 50ml Falcon tube.
  2. Add 50μl RNase A (20mg/ml) and incubate at room temperature for 30 min.
  3. Add 5ml phenol, and shake vigorously for 1 minute, spin down at maximum speed in the swing out rotor for 15 minutes.
  4. Slowly recover the water phase (upper) and make sure you leave the interphase in the tube. Transfer to a new tube and add 5ml of chloroform, shake again as before and spin as before for 5 minutes.
  5. Slowly recover the water phase (this time it is easier to avoid the interphase), transfer to a 15ml Corex tube (approx 5ml), add 500μl 5M NaClO4 (10% of the water volume), mix and add 4ml isopropanol (80% of the water volume). Seal the tube with parafilm, and turn it around a few times to mix everything.
  6. Then spin down in HB4 rotor at 10,000rpm for 15 minutes.
  7. Discard the supernatant, dry the pellet, resuspend in 500 µl of TE (10mM Tris-HCl pH 8.0; 0.1 mM EDTA pH 8.0) and transfer to a sterile eppendorf tube.
  8. Store at 4°C.
  9. Check the DNA on an agarose gel to make sure most of the RNA is gone. Determine the DNA concentration and purity spectrophotometrically (a nanodrop is a useful tool at this point).

Clean Up of DNA

The reaction conditions of many recombinant DNA procedures are incompatible, for example, between a restriction digest and a ligation. This requires that you remove the enzyme and buffer from the previous step so that you can start the next step. This can be done through various methods including using extraction and purification kits, phenol-chloroform extraction, or using miniprep kits with a modified method. Probably the most important consideration for most labs is cost. A phenol-chloroform extraction is the cheapest and using a miniprep spin column being the most expensive. However, in terms of safety this is the exact opposite!

Many specialist kits exist for cleaning up reactions. Many people use PCR clean up kits , such as the GenElute™ PCR Cleanup Kit (Catalog No. NA1020) for the cleaning up of enzymatic reactions but do check the size exclusion limits on the column and the binding capacities. Some PCR kits don’t bind DNA <200bp or >4-5kb.

Below is method for using Phenol and Chloroform to extract DNA from an enzymatic reaction but read the SDS before proceeding. The steps to recover the aqueous upper-phase are a bit tricky, perhaps the most difficult of all recombinant DNA techniques, and it is worthwhile training it on fake samples (just TE) before you start. Shaky hands are not helpful.

Reagents

  • TE (10mM Tris-HCl pH 8.0, 0.1mM EDTA)
  • Equilibrated Phenol
  • Chroloform
  • 5M Sodium perchlorate (5M NaClO4)

Potential Hazard: Phenol is very dangerous and causes skin burns immediately on contact. It is toxic on contact, by inhalation and by consumption. Read and understand the SDS before using any hazardous chemicals.

Potential Hazard: Chloroform is toxic by inhalation, oral consumption and skin contact. Read and understand the SDS before using any hazardous chemicals.

  1. Place 50μl of “dirty” DNA in a 1.5ml eppendorf tube.
  2. Add 50 μl of TE.
  3. Add 50 μl of equilibrated phenol.
  4. Vortex (the sample should become “milky”). The idea is to get an emulsion between the phenol and the water phase. Be careful to ensure the lid is properly shut. Proteins will migrate to the phenol phase, whereas the DNA will stay in the water phase.
  5. Add 100 μl of chloroform and vortex. Be careful to ensure the lid is properly shut. Now that the DNA and proteins are partitioned, the idea is to separate the two phases.
  6. Centrifuge the tube for 5 minutes at maximum speed (12-13000 rpm) in a microcentrifuge.
  7. Recover the upper (aqueous) phase and transfer to a new 1.5 ml tube containing 100 μl of chloroform.
  8. Vortex and centrifuge again for 2 minutes at maximum speed in a microcentrifuge.
  9. Recover the upper (aqueous) phase in a new 1.5 ml tube and then add 10 μl of 5M NaClO4 and mix.
  10. Add 110 μl of Isopropanol, vortex and centrifuge 15 minutes at maximum speed in a microcentrifuge.
  11. Carefully discard the supernatant and centrifuge again for 1 minute at maximum speed.
  12. Insert either a fine pipette tip or fine glass pasteur pipette to the opposite side of the pellet (which should be barely visible) and suck out the liquid. Keep sucking even when the liquid has gone, to remove all liquid from the walls of the tube. You should also try to collect all the droplets that you see on the walls of the tube by moving the pipette and using the capillary forces. It is important to have only a dry pellet left.
  13. Leave the tube open for 1 minute at room temperature and then resuspend the pellet it in an appropriate volume of TE (approx 30-50μl).
  14. Keep the tube on ice and vortex now and then, it does take a few minutes before all of the plasmid in the pellet is dissolved again.
  15. Now you can move to the next manipulation, for example a ligation.

Tip: Disposing of phenol waste can be difficult in many labs. Ideally, contact the person in charge of safety and disposals in your building and ask them what you need to do so that you comply with local regulations.