Team:Amsterdam/Labwork/Protocols

From 2011.igem.org

Revision as of 15:39, 21 June 2011 by Pvandieken (Talk | contribs)

Contents

Making LB Medium (P1.)

Luria-Bertani Medium (aka L-Broth or LB Medium). (Bertani says LB really stands for lysogeny broth.) LB is a standard growth medium for a variety of bacteria and conditions.

Ingredients

  1. 10 g Bacto-tryptone
  2. 5 g yeast extract
  3. 10 g NaCl

Note: There are two formulations of LB, Miller and Lennox, that differ in the amount of NaCl. Lennox has less salt, only 5 g/L. The Qiagen miniprep kit recommends LB with 10 g NaCl for highest plasmid yields.

Protocol

  1. Mix dry ingredients and add distilled water up to 1 Liter
  2. Pour into 2 L flask (or greater)
  3. Autoclave (liquid cycle)
    • 250°F, 22psi, 30 minutes

Notes: We do not pH medium when we make it on the fly. However, if it is really important, pH the medium to 7.0 with 5M NaOH (~200µL). We usually obtain this from the kitchen.

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

Making SOB Medium (P2.)

SOB Medium. Used in growing bacteria for preparing chemically competent cells

Ingredients

  • 0.5% (w/v) yeast extract
  • 2% (w/v) tryptone
  • 10 mM NaCl
  • 2.5 mM KCl
  • 20 mM MgSO4

Per liter:

  • 5 g yeast extract
  • 20 g tryptone
  • 0.584 g NaCl
  • 0.186 g KCl
  • 2.4 g MgSO4

Note: Some formulations of SOB use 10 mM MgCl2 and 10 mM MgSO4 instead of 20 mM MgSO4.

SOB medium is also available dry premixed from Difco, 0443-17.

Adjust to pH 7.5 prior to use. This requires approximately 25 ml of 1M NaOH per liter.

15/10 medium

Growth of competent TOP10 cells in Example 2 of the Bloom05 patent is performed in 15/10 broth, which is similar to SOB:

  • 1.5% yeast extract
  • 1% Bacto-Tryptone
  • 10mM NaCl
  • 2mM KCl
  • 10 mM MgCl2
  • 10 mM MgSO4

Source

Adapted From:

F. Ausubel et al., Short Protocols in Molecular Biology (John Wiley & Sons, ed. 4, 1999) pg. A1-36

Making SOC Medium (P3.)

SOC Medium.

Ingredients

  • SOB
  • 20 mM glucose

Protocol

  1. Follow directions to make 1 liter of Making SOB Medium (P2.) media
  2. After cooling medium to less than 50°C, add 20 ml filter sterilized 20% glucose solution

Source

Adapted From:

F. Ausubel et al., Short Protocols in Molecular Biology (John Wiley & Sons, ed. 4, 1999) pg. A1-36

Making Competent Cells (P4.)

This protocol is a variant of the Hanahan protocol Hanahan91 using CCMB80 buffer for DH10B, TOP10 and MachI strains. It builds on Example 2 of the [http://openwetware.org/images/b/bd/Pat6855494.pdf Bloom05 patent] as well. This protocol has been tested on NEB10, TOP10, MachI and [http://openwetware.org/wiki/Talk:TOP10_chemically_competent_cells BL21(DE3)] cells. See [http://openwetware.org/wiki/Bacterial_Transformation OWW Bacterial Transformation page] for a more general discussion of other techniques. The [http://openwetware.org/images/0/0c/Pat6960464.pdf Jesse '464 patent] describes using this buffer for DH5α cells. The [http://openwetware.org/images/c/c2/Pat6709852.pdf Bloom04] patent describes the use of essentially the same protocol for the Invitrogen Mach 1 cells.

This is the chemical transformation protocol used by Tom Knight and the [http://partsregistry.org Registry of Standard Biological Parts].

Materials

  • Detergent-free, sterile glassware and plasticware (see procedure)
  • Table-top OD600nm spectrophotometer
  • SOB

CCMB80 buffer

  • 10 mM KOAc pH 7.0 (10 ml of a 1M stock/L)
  • 80 mM CaCl2.2H2O (11.8 g/L)
  • 20 mM MnCl2.4H2O (4.0 g/L)
  • 10 mM MgCl2.6H2O (2.0 g/L)
  • 10% glycerol (100 ml/L)
  • adjust pH DOWN to 6.4 with 0.1N HCl if necessary
    • adjusting pH up will precipitate manganese dioxide from Mn containing solutions.
  • sterile filter and store at 4°C
  • slight dark precipitate appears not to affect its function

Procedure

Preparing glassware and media

Eliminating detergent

Detergent is a major inhibitor of competent cell growth and transformation. Glass and plastic must be detergent free for these protocols. The easiest way to do this is to avoid washing glassware, and simply rinse it out. Autoclaving glassware filled 3/4 with DI water is an effective way to remove most detergent residue. Media and buffers should be prepared in detergent free glassware and cultures grown up in detergent free glassware.

Prechill plasticware and glassware

Prechill 250mL centrifuge tubes and screw cap tubes before use.

Preparing seed stocks

  • Streak TOP10 cells on an SOB plate and grow for single colonies at 23°C
    • room temperature works well
  • Pick single colonies into 2 ml of SOB medium and shake overnight at 23°C
    • room temperature works well
  • Add glycerol to 15%
  • Aliquot 1 ml samples to Nunc cryotubes
  • Place tubes into a zip lock bag, immerse bag into a dry ice/ethanol bath for 5 minutes
    • This step may not be necessary
  • Place in -80°C freezer indefinitely.

Preparing competent cells

  • Inoculate 250 ml of SOB medium with 1 ml vial of seed stock and grow at 20°C to an OD600nm of 0.3
    • This takes approximately 16 hours.
    • Controlling the temperature makes this a more reproducible process, but is not essential.
    • Room temperature will work. You can adjust this temperature somewhat to fit your schedule
    • Aim for lower, not higher OD if you can't hit this mark
  • Centrifuge at 3000g at 4°C for 10 minutes in a flat bottom centrifuge bottle.
    • Flat bottom centrifuge tubes make the fragile cells much easier to resuspend
    • It is often easier to resuspend pellets by mixing before adding large amounts of buffer
  • Gently resuspend in 80 ml of ice cold CCMB80 buffer
    • sometimes this is less than completely gentle. It still works.
  • Incubate on ice 20 minutes
  • Centrifuge again at 4°C and resuspend in 10 ml of ice cold CCMB80 buffer.
  • Test OD of a mixture of 200 μl SOC and 50 μl of the resuspended cells.
  • Add chilled CCMB80 to yield a final OD of 1.0-1.5 in this test.
  • Incubate on ice for 20 minutes
  • Aliquot to chilled screw top 2 ml vials or 50 μl into chilled microtiter plates
  • Store at -80°C indefinitely.
    • Flash freezing does not appear to be necessary
  • Test competence (see below)
  • Thawing and refreezing partially used cell aliquots dramatically reduces transformation efficiency by about 3x the first time, and about 6x total after several freeze/thaw cycles.

Measurement of competence

  • Transform 50 μl of cells with 1 μl of standard pUC19 plasmid (Invitrogen)
    • This is at 10 pg/μl or 10-5 μg/μl
    • This can be made by diluting 1 μl of NEB pUC19 plasmid (1 μg/μl, NEB part number N3401S) into 100 ml of TE
  • Hold on ice 0.5 hours
  • Heat shock 60 sec at 42C
  • Add 250 μl SOC
  • Incubate at 37 C for 1 hour in 2 ml centrifuge tubes rotated
    • using 2ml centrifuge tubes for transformation and regrowth works well because the small volumes flow well when rotated, increasing aeration.
    • For our plasmids (pSB1AC3, pSB1AT3) which are chloramphenicol and tetracycline resistant, we find growing for 2 hours yields many more colonies
    • Ampicillin and kanamycin appear to do fine with 1 hour growth
  • Plate 20 μl on AMP plates using sterile 3.5 mm glass beads
    • Good cells should yield around 100 - 400 colonies
    • Transformation efficiency is (dilution factor=15) x colony count x 105/µgDNA
    • We expect that the transformation efficiency should be between 5x108 and 5x109 cfu/µgDNA

5x Ligation Adjustment Buffer

  • Intended to be mixed with ligation reactions to adjust buffer composition to be near the CCMB80 buffer
  • KOAc 40 mM (40 ml/liter of 1 M KOAc solution, pH 7.0)
  • CaCl2 400 mM (200 ml/l of a 2 M solution)
  • MnCl2 100 mM (100 ml/l of a 1 M solution)
  • Glycerol 46.8% (468 ml/liter)
  • pH adjustment with 2.3% of a 10% acetic acid solution (12.8ml/liter)
    • Previous protocol indicated amount of acetic acid added should be 23 ml/liter but that amount was found to be 2X too much per tests on 1.23.07 --Meagan 15:50, 25 January 2007 (EST)
  • water to 1 liter
  • autoclave or sterile filter
  • Test pH adjustment by mixing 4 parts ligation buffer + 1 part 5x ligation adjustment buffer and checking pH to be 6.3 - 6.5
  • Reshma P. Shetty 10:49, 11 February 2008 (CST): Use of the ligation adjustment buffer is optional.

References

  1. Hanahan91 pmid=1943786
  2. Reusch86 pmid=3536850
  3. Addison04 pmid=15470891
  4. Bloom04 US Patent 6,709,852 [http://openwetware.org/images/c/c2/Pat6709852.pdf pat6709852.pdf]
  5. Bloom05 US Patent 6,855,494 [http://openwetware.org/images/b/bd/Pat6855494.pdf pat6855494.pdf]
  6. Jesse05 US Patent 6,960,464 [http://openwetware.org/images/0/0c/Pat6960464.pdf pat6960464.pdf]

Linearized Plasmid Backbones (P4.)

This protocol was developed by Tom Knight, samples of standard Registry plasmid backbones prepared using this method were sent out in the Spring 2011 DNA Distribution kits.

Short single stranded DNA fragments will not ligate to 4 bp overhangs. By creating a very short overhang on a PCR of a plasmid backbone, the remnant, when cut with EcoRI and PstI is sufficiently short that it will not anneal at ligation temperature, and will therefore not ligate. This allows us to build high quality construction plasmid backbone without purifying away the cut fragments remaining after PCR.

We are distributing the prepared construction plasmid as purified PCR products, diluted to standard concentration, but prior to cutting with EcoRI and PstI. Standard assembly will cut this plasmid backbone with EcoRI and PstI at the same time that the two assembled fragments are cut with EcoRI and SpeI and with XbaI and PstI, respectively.

The preparation of this PCR fragment is done with primers having short overhangs past the EcoRI and PstI sites, followed by PCR cleanup, dilution to standard concentration, and quality control testing.

Note: The Registry shipping plasmid backbone is pSB1C3. If you are making linearized plasmid backbone in order to send parts to the Registry, you must use pBS1C3.


Primers: 

gccgctgcagtccggcaaaaaa,SB-prep-3P-1
atgaattccagaaatcatccttagcg,SB-prep-2Ea

Diluted to 30 pmol/ul

These primers have been tested with pSB1C3, pSB1A3, pSB1K3, and pSB1T3.

Using the Linearized Plasmid Backbones

The Spring 2011 DNA Distribution should come with a set of linearized plasmid backbones: pSB1A3, pSB1C3, pSB1K3, and pSB1T3. The linearized plasmid backbones (25ng/ul at 50ul) should be stored at 4C or lower. Prior to ligation the plasmid backbones need to be cut with EcoRI and PstI.

Digest

  • Enzyme Master Mix for Plasmid Backbone (25ul total, for 6 rxns)
    • 5 ul NEB Buffer 2
    • 0.5 ul BSA
    • 0.5 ul [http://www.neb.com/nebecomm/products/productR3101.asp EcoRI-HF]
    • 0.5 ul [http://www.neb.com/nebecomm/products/productR0140.asp PstI]
    • 0.5 ul [http://www.neb.com/nebecomm/products/productR0176.asp DpnI]
    • 18 ul dH20
  • Digest Plasmid Backbone
    • Add 4 ul linearized plasmid backbone (25ng/ul for 100ng total)
    • Add 4 ul of Enzyme Master Mix
    • Digest 37C/30 min, heat kill 80C/20 min


Ligation

  • Add 2ul of digested plasmid backbone (25 ng)
  • Add equimolar amount of EcoRI-HF SpeI digested fragment (< 3 ul)
  • Add equimolar amount of XbaI PstI digested fragment (< 3 ul)
  • Add 1 ul [http://www.neb.com/nebecomm/products/productm0202.asp T4 DNA ligase buffer]. Note: Do not use quick ligase
  • Add 0.5 ul [http://www.neb.com/nebecomm/products/productm0202.asp T4 DNA ligase]
  • Add water to 10 ul
  • Ligate 16C/30 min, heat kill 80C/20 min
  • Transform with 1-2 ul of product

Making Linearized Plasmid Backbones (P5.)

Bulk Production

The following is the protocol that we used to create the linearized plasmid backbones shipped with the Spring 2011 DNA Distribution. The protocol is in 96 well format, but may be scaled down to suit smaller batches.

2011 Plasmid Backbone Production

PCR mix

  • 9.6ml of [http://products.invitrogen.com/ivgn/product/10790020?ICID=search-10790020 PCR Supermix High Fidelity]
  • 67 ul of primer SB-prep-2Eb
  • 67 ul of primer SB-prep-3P-1
  • 10 ul of template DNA at 10ng/ul (100ng total) (Note: Do not use a sample of linearized plasmid backbones (PCRed) as a template)
  • Aliquot 100ul per well in 96 well plate

PCR program

  1. 95C/2min
  2. 95C/30s
  3. 55C/30s
  4. 68C/3min
  5. Repeat cycle (steps 2 to 4, 37 more times)
  6. 68C/10min

PCR cleanup

Purification of 96 well plates was done through Promega [http://www.promega.com/products/dna-and-rna-purification/dna-fragment-purification/wizard-sv-96-pcr-clean_up-system/ Wizard SV 96 PCR Clean-Up kit] and a vacuum manifold. The protocol below follows the manual, with a few changes (in bold), however please see manual for setup instructions.

  1. Add equal volume of Binding Solution to PCR product (add 100ul of Binding Solution to 100ul of product)
  2. Mix by pipetting, transfer all 200ul to Binding Plate, let sit for 1 min
  3. Apply vacuum until samples pass through, about 30s to 1 min
  4. Add 200 ul of freshly prepared 80% ethanol to Binding Plate, let sit for 1min, apply vacuum until ethanol passes through, about 20s to 1 min.
  5. Repeat ethanol wash (step 4) twice more for three washes total
  6. Remove Binding Plate from wash manifold, blot on kim wipes, reinstall in wash manifold
  7. Apply vacuum for 4 min to fully dry Binding Plate
  8. Remove Binding Plate from wash manifold, blot on kim wipes, reinstall in collection manifold
  9. Add 50ul of TE buffer, let sit for 1 min, apply vacuum until eluted, about 1 min
  10. Repeat 50ul elution (step 9) for a total elution of 100ul
  11. Measure concentration on nanodrop, adjust to 25 ng/ul with TE


Single Reaction PCR

PCR mix

  • 100 ul [http://products.invitrogen.com/ivgn/product/10790020?ICID=search-10790020 PCR Supermix High Fidelity]
  • 0.7 ul of SB-prep-3P-1
  • 0.7 ul of SB-prep-2Ea
  • 0.5 ul template DNA at 10 ng/ul (Note: Do not use a sample of linearized plasmid backbones (PCRed) as a template)

PCR program

  1. 94C/2min
  2. 94C/30s
  3. 55C/30s
  4. 68C/3min
  5. Repeat cycle (steps 2 to 4, 35 more times)
  6. 68C/10min
  7. Digest with DpnI enzyme: 2ul in 100ul reaction, incubate 37C/hour; heat kill 80C/20min

PCR cleanup

[http://www.qiagen.com/products/dnacleanup/gelpcrsicleanupsystems/qiaquickpcrpurificationkit.aspx QIAquick PCR Purification]

  • Add 500 ul Qiagen buffer PB
  • Spin through a column twice, discard flowthrough
  • Wash 1x with 700 ul buffer PB
  • Wash 2x with 760 ul buffer PE
  • Discard liquid, spin dry at 17000g for 3 min
  • Elute into a new tube twice with 50 ul of TE (100 ul total)


Quality Control

We recommend QCing constructed linearized plasmid backbones, to test success of PCR, ligation efficiency, and background.

  1. Run unpurified PCR product (1 ul) on a gel to verify the correct band and concentration and lack of side products.
  2. Test concentration of purified PCR product. Note: Expected yield should be 40ng/ul or higher. Adjust to 25ng/ul with TE.
  3. Run a digest and ligation test with purified PCR product to determine EcoRI and PstI cutting and ligation efficiency.


Digest

  • Digest Master Mix (10rxns)
    • 15 ul NEB Buffer 2
    • 1.5 ul BSA
    • 90 ul dH20
  • Run Digest
    • 4 ul of plasmid backbone (approximately 100 ng)
    • 10.5 ul of Digest Master Mix
    • 0.5 ul either EcoRI-HF or PstI enzyme (not both!)
    • Digest 37C/30min; 80C/20 min
    • Proceed directly to ligation


Ligation

  • Ligation Master Mix (10rxns)
    • 20 ul T4 DNA ligase buffer
    • 5 ul T4 DNA ligase
    • 25 ul water
  • Ligation Test
    • Add 5 ul of ligation master mix to digested product
    • Ligate 16C/30min; 80C/20 min
    • Run all 20 ul on a gel
    • Compare intensity of the single and double length bands. More efficient ligations will show stronger double length bands than single.


Transformation test

  • Transform 1 ul of the diluted final product into highly competent cells
  • Control transform 10 pg of pUC19
  • Plate on the appropriate antibiotic
  • Observe few colonies. Any colonies represent background to the three antibiotic assembly process
  • Quantify the effective amount of remaining circular DNA able to transform

Transforming Competent Cells (P7.)

Estimated time: 3 hours (plus 12-14 hour incubation)
It is important to note that we have tested transformations of the distribution kit with this protocol. We have found that it is the best protocol to use with BioBrick parts and ensures the highest efficiency for the transformation. This protocol may be particularly useful if you are finding that your transformations are not working, or yielding few colonies.


Materials needed

  • Resuspended DNA (Resuspend well in 10ul dH20, pipette up and down several times, let sit for a few minutes)
  • Competent cells (50ul per a transformation)
  • Ice
  • 42º water bath
  • 37º incubator
  • SOC (check for contamination!)
  • Petri dishes with LB agar and appropriate antibiotic (two per a transformation)

Protocol

  1. Start thawing the competent cells on crushed ice.
  2. Add 50 µL of thawed competent cells and then 1 - 2 µL of the resuspended DNA to the labelled tubes. Make sure to keep the competent cells on ice.
  3. Incubate the cells on ice for 30 minutes.
  4. Heat shock the cells by immersion in a pre-heated water bath at 42ºC for 60 seconds. A water bath improves heat transfer to the cells.
  5. Incubate the cells on ice for 5 minutes.
  6. Add 200 μl of SOC broth (make sure that the broth does not contain antibiotics and is not contaminated)
  7. Incubate the cells at 37ºC for 2 hours while the tubes are rotating or shaking. Important: 2 hour recovery time helps in transformation efficiency, especially for plasmids with antibiotic resistance other than ampicillin.
  8. Label two petri dishes with LB agar and the appropriate antibiotic(s) with the part number, plasmid, and antibiotic resistance. Plate 20 µl and 200 µl of the transformation onto the dishes, and spread. This helps ensure that you will be able to pick out a single colony.
  9. Incubate the plate at 37ºC for 12-14 hours, making sure the agar side of the plate is up. If incubated for too long the antibiotics start to break down and un-transformed cells will begin to grow. This is especially true for ampicillin - because the resistance enzyme is excreted by the bacteria, and inactivate the antibiotic outside of the bacteria.

Miniprepping (P8.)

After you have transformed a part from one of our distributions you will want to miniprep it so we have provided the protocol that is used at iGEM HQ. Here at iGEM we use [http://www1.qiagen.com/Products/Plasmid/QIAprepMiniprepSystem/QIAprepSpinMiniprepKit.aspx Qiagen Spin Miniprep Kits] for doing small batches of minipreps.


Protocol

See [http://www1.qiagen.com/literature/handbooks/PDF/PlasmidDNAPurification/PLS_QP_Miniprep/1027678_HB_QP_0504_WW_LR.pdf here] or [http://www1.qiagen.com/literature/protocols/QIAprepMiniprep.aspx here] for the handbook for the Qiagen Spin Miniprep Kit. If you have never done this protocol before, read the the background information in the handbook (like the Important Notes section). It contains useful information. The following has been reproduced from the handbook and annotated based on experience with the kit.

Protocol: QIAprep Spin Miniprep Kit Using a Microcentrifuge

This protocol is designed for purification of up to 20 μg of high-copy plasmid DNA from 1–5 ml overnight cultures of E. coli in LB (Luria-Bertani) medium. For purification of low-copy plasmids and cosmids, large plasmids (>10 kb), and DNA prepared using other methods, refer to the recommendations on page 37. Please read “Important Notes” on pages 19–21 before starting. Note: All protocol steps should be carried out at room temperature.

Procedure

  1. Resuspend pelleted bacterial cells in 250 µl Buffer P1 (kept at 4 °C) and transfer to a microcentrifuge tube.
    Ensure that RNase A has been added to Buffer P1. No cell clumps should be visible after resuspension of the pellet.
  2. Add 250 μl Buffer P2 and gently invert the tube 4–6 times to mix.
    Mix gently by inverting the tube. Do not vortex, as this will result in shearing of genomic DNA. If necessary, continue inverting the tube until the solution becomes viscous and slightly clear. Do not allow the lysis reaction to proceed for more than 5 min.
  3. Add 350 μl Buffer N3 and invert the tube immediately but gently 4–6 times.
    To avoid localized precipitation, mix the solution gently but thoroughly, immediately after addition of Buffer N3. The solution should become cloudy.
  4. Centrifuge for 10 min at 13,000 rpm (~17,900 x g) in a table-top microcentrifuge.
    A compact white pellet will form.
  5. Apply the supernatants from step 4 to the QIAprep spin column by decanting or pipetting.
  6. Centrifuge for 30–60 s. Discard the flow-through.
    Spinning for 60 seconds produces good results.
  7. (Optional): Wash the QIAprep spin column by adding 0.5 ml Buffer PB and centrifuging for 30–60 s. Discard the flow-through.
    This step is necessary to remove trace nuclease activity when using endA+ strains such as the JM series, HB101 and its derivatives, or any wild-type strain, which have high levels of nuclease activity or high carbohydrate content. Host strains such as XL-1 Blue and DH5α™ do not require this additional wash step.
    Although they call this step optional, it does not really hurt your yield and you may think you are working with an endA- strain when in reality you are not. Again for this step, spinning for 60 seconds produces good results.
  8. Wash QIAprep spin column by adding 0.75 ml Buffer PE and centrifuging for 30–60 s.
    Spinning for 60 seconds produces good results.
  9. Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer.
    IMPORTANT: Residual wash buffer will not be completely removed unless the flow-through is discarded before this additional centrifugation. Residual ethanol from Buffer PE may inhibit subsequent enzymatic reactions. They are right about this.
  10. Place the QIAprep column in a clean 1.5 ml microcentrifuge tube. To elute DNA, add 50 μl Buffer EB (10 mM Tris·Cl, pH 8.5) or water to the center of each QIAprep spin column, let stand for 1 min, and centrifuge for 1 min.
    If you are concerned about the concentration of the DNA, you can alternatively add 30 μL water to the center of the column, incubate at room temperature on the bench for 5 mins and then centrifuge for 1 min. This will increase the concentration of DNA in your final sample which can be useful in some cases. See notes below for why you should elute in water rather than the Buffer EB they recommend if you plan to sequence your sample. Even if you are not sequencing, it may be beneficial to elute in water. For instance, if you elute in buffer EB and you are using this DNA in a restriction digest, then the additional salts in your sample can affect the salt content of your digest. This may matter with some finicky enzymes.

Notes

  • Heating the elution buffer to 55°C prior to loading on the column can slightly increase yields.
  • Similarly, doing the elution in two steps (first a 30 μL elution and then a 20 μL dilution) can also slightly increase yields.


Materials

Do not autoclave solutions containing isopropanol or MOPS; use sterile filtration if necessary.

Buffer P1

  • 50 mM Tris-HCl pH 8.0
  • 10 mM EDTA
  • 100 μg/ml RNaseA

The buffer and RNaseA can also be ordered from Qiagen separately (catalog numbers 19051 and 19101).

Buffer P2

  • 200 mM NaOH
  • 1% SDS

Buffer P3 (not for spin columns, but for Qiatips, midi, maxi, giga kits)

  • 3.0 M potassium acetate pH 5.5

Buffer N3

  • 4.2 M Gu-HCl
  • 0.9 M potassium acetate
  • pH 4.8

Buffer PB

  • 5 M Gu-HCl
  • 30% ethanol
  • (maybe add 10mM Tris-HCL PH 6.6, and that is better)

Buffer PE

  • 10 mM Tris-HCl pH 7.5
  • 80% ethanol

Buffer QBT equilibration buffer

  • 750 mM NaCl
  • 50 mM MOPS pH 7.0
  • 15% isopropanol
  • 0.15% triton X-100

Buffer QC wash buffer

  • 1.0M NaCl
  • 50 mM MOPS pH 7.0
  • 15% isopropanol

Buffer QF elution buffer

  • 1.25M NaCl
  • 50 mM Tris-HCl pH 8.5
  • 15% isopropanol

Buffer QN

  • 1.6M NaCl
  • 50 mM MOPS pH 7.0
  • 15% isopropanol


Buffer FWB2

  • 1M potassium acetate, pH 5.0


(Source: [http://methodsandreagents.pbwiki.com/], US Patent 6,383,393)

The LyseBlue indicator dye added to some of the buffers is Thymophthalein, pH shift from colorless to blue at pH 9.3

Restriction Digest (P9.)

At iGEM HQ we use this protocol for restriction digests along with enzymes purchased from NEB.

Materials

  • PCR tube
  • dH20
  • Enzymes (EcoRI, XbaI, SpeI, PstI)
  • BSA
  • Enzyme Buffer (NEBuffer 2)*

Notes: You should keep all materials on ice.

Protocol

1. Add 500ng of DNA to be digested, and adjust with dH20 for a total volume of 42.5ul.
2. Add 5ul of NEBuffer 2 to the tube.
3. Add 0.5ul of BSA to the tube.
4. Add 1ul of your first enzyme.
5. Add 1ul of your second enzyme.
6. There should be a total volume of 50ul. Mix well and spin down.
7. Incubate the restriction digest at 37C for 30min, and then 80C for 20min to heat kill the enzymes.
We incubate in a thermocycler with a heated lid
8. Run a portion of the digest on a gel, to check that both plasmid and part length are accurate. You may also use 2ul of the digest (20ng of DNA) for ligations.

Ligations (P10.)

After following our restriction digest protocol (which uses 500ng of DNA) you may follow these steps for ligation.

1. Add 11ul of dH20
2. Add 2ul from each sample you will be ligating (destination plasmid, and part)
3. Add 2ul of T4 DNA Ligase Reaction Buffer
4. Add 1ul of T4 DNA Ligase
5. Mix well, and spin down.
6. Incubate for 30min at 16C and 20min at 80C to heat kill.
7. Use 2ul of ligation to transform into competent cells.

Retrieved from "http://2011.igem.org/Team:Amsterdam/Labwork/Protocols"