Difference between revisions of "Team:MIT/Protocols"

This is the media used for C. Hutch and co cultures with E. Coli. The media contains all the salts needed for both bacteria but not the carbon source (filter paper) due to the carbon source not being liquid. Cultures are made by adding the salt solution to a tube containing a piece of filter paper where the tube with the paper in it has been sterilized (usually by autoclave).

C Hutch ALONE in 1 L

Paper: 50 grams per liter of media, which is one 0.5g 9cm No. 1 filter paper disks per 100 mL.

Autoclave 15 min.

E Coli ALONE (M9) in 1 L

1. Make 5x M9 salts
   • To make M9 Salts aliquot 800ml H₂O and add
     • 64g Na₂HPO₄-7H₂O
     • 15g KH₂PO₄
     • 2.5g NaCl
     • 5.0g NH₄Cl
     • Stir until dissolved
     • Adjust to 1000ml with distilled H₂O
     • Sterilize by autoclaving
2. Measure ~700ml of distilled H₂O (sterile)
3. Add 200ml of M9 salts
4. Add 2ml of 1M MgSO₄ (sterile)
5. Add 20 ml of 20% glucose (or other carbon source)
6. Add 100ul of 1M CaCl₂ (sterile)
7. Adjust to 1000ml with distilled H₂O

C Hutch is slow to grow, so experiments will be much faster if large initial concentrations of C Hutch is used. This is achieved by having a continual stock of C Hutch in the incubator which is used to provide cells for experiments

Protocol:

Stripette:

• 10ml of current stock
• 90ml of DSM3T media
• 5g of autoclaved filter paper strips

into a 500ml flask, and place in shaker.

1. Make sure that the incubator (30/37C) and heat block (42C) are ON.
   1. Put water in the wells of the 42°C heat block.
2. Make sure required antibiotic plates are present.  Make sure you're using the right antibiotic plates for your plasmid's resistance!
   1. Warm plates to 37°C.  Cold plates reduce transformation efficiency by an order of magnitude.
   2. Also warm 500 µl SOC per transformation to room temperature (if it was in the refrigerator.)
3. Take the DNA out of --20 freezer, let it thaw.
   1. Vortex DNA to mix, then spin down.  Make sure it is completely thawed out!
4. Make sure that all of the required reagents/DNA etc are present at the site of transformation before you take the cells out of the -80.
5. Thaw the competent cells on ice for 3-4 min.
   1. You want to add your DNA right as the last bit of cells' ice melts.  Even if it's still a little slushy, that's okay.
6. Add 1-2 µl of DNA into the comp cells.  Stir with a pipette tip a few times, then put right back on ice.
   1. If you're transforming the result of a reaction (GG, LR, etc) add 1-2 µl of the reaction.  Don't add more: many of these reactions have additives that screws up transformation.
   2. If you're transforming plasmid DNA (from a miniprep), either (a) dilute it out so you add only ~10 ng of DNA, or (b) plate only 10 µl of the outgrowth – else you'll get a lawn!  Super-coiled DNA transforms super-efficiently. As an alternative, transform 1 ul of miniprep DNA and then streak the outgrowth instead of plating with beads.
   3. You can rescue DNA from an empty mini prep tube by gently pipetting the cells into the empty DNA tube, enough DNA will be stuck to the walls.
7. Incubate the cells on ice for 30-40 min.
8. Heat shock the cells for EXACTLY 30 sec at 42 C water bath.
9. Place back on ice for 90 seconds.
10. Add 250 ul of SOC (37° to RT) medium to each tube (S.O.C is made by dissolving 0.5 ml of 20% glucose in 25 ml of SOB. Make sure that the SOC is clear and not cloudy/ contaminated.)
11. Shake the tubes at 37 C, 280 rpm for 60 min.
12. Plate 100 µl for a reaction product, or 10 µl in a 100 µl puddle of water for a supercoiled plasmid.
13. Incubate plates upside down overnight at 37 C or 16-18h at 30C.
    Can leave the cells in the incubator for up to 18 hours but no more.

Preparing the Gel

1. Check to see if there is a gel waiting in the fridge.
2. Dissolve UltraPure agarose to a final concentration of 1%(by mass) in TAE buffer in a glass bottle.
3. Heat the solution in the microwave with frequent stirring to dissolve the agarose homogenously. ~1 minute/200ml solution
4. Let sit until cool enough to handle.
5. Add 10 µl SYBRSafe (1:10000) per 100 ml of the solution and mix well.
6. Pour 50ml* of solution per small gel tray. The gel trays and combs should be pre-cleaned with water and wiped dry.
   • Note for combs: 15-well combs hold about 6 ul liquid per well, 12-well combs hold about 15 ul per well, 8-well combs hold about 20 ul per well
     • Taping two 8-well comb wells together results in a well that holds up to 100 ul
     • Taping three 8-well comb wells together result in a well that holds up to 200 ul
7. Use 120 ml per large gel tray. [need to update amounts]
   1. For the small set: small trays hold 20 ml, large trays hold 50 ml
8. Wait for the gels to solidify. ~15 mins
9. Label and store at 4C.

*Most of the iGEM gel trays are the small trays that fit 50ml of solution for making the gel.

Running the Gel

When doing gel extraction, it is important to run both an analytical gel (to view under UV) and an extraction gel (from which bands are excised). UV damages DNA, and so we dont want to expose our extracted DNA.

Analytical Gel:

The analytical gel should have between 20 and 100 ng of DNA in each well. It should be an exact copy of the extraction gel with respect to position, voltage, and run time.

Extraction Gel:

This should be the rest of the digestion(s).

The analytical and extraction gels can technically be part of the same physical gel. Make sure to separate with a razor blade before imaging.

Refer to Gel Prep protocol above to determine the amounts of liquid to load for the specific well.

Appropriate Hyperladder to be used for PCR product which is linear. Usually Hyperladder I will be used.

1. While casting gel, add two sets of lanes; use one set to load an analytical gel. 
2. Add 2ul gel loading buffer (Orange G 6X; it helps DNA sink into the bottom of the well) to DNA.
3. Make sure there is enough 1xTAE in the plate holder.
4. Load 5.0ul of appropriate hyperladder to one of the lanes.
5. Load appropriate amount of DNA - As much as possible! Usually 15-18ul - (mixed with the buffer) in each well.
6. Set the timer and voltage to 100V and 25 min.

Analytical Gel Annotation

The following things need to be added to the analytical gel image BEFORE it is posted to the wiki:

• Label each lane with part number and amount of DNA loaded
• Label each band with length and proposed identification
• Include wt% agarose, run time, and voltage

Gel Extraction Protocol using Zymo kit (preferred if available)

1. Place the extraction gel on the blue light table.
2. Cut out the appropriate bands. Place into 2mL microtube(s). Try to cut out as small a piece as possible while still getting all the DNA.
3. Weigh gel slice (tare with empty microtube). Add 3 volumes of ADB buffer per mg of gel (so a 100mg gel gets 300 uL of ADB buffer).
4. Incubate at 55C for 10 minutes. Make sure that the gel is completely dissolved.
5. Add dissolved gel solution to Zymo column in collection tube. Max volume is 800 uL at a time.
6. Spin 14000 rpm for 30 sec.
7. Discard liquid in collection tube.
8. Repeat step 5-7 if had more than 800 uL dissolve gel.
9. Add 200 uL DNA wash buffer.
10. Spin 14000 rpm 30 seconds.
11. Discard liquid in collection tube.
12. Add 200 uL DNA wash buffer
13. Spin 14000 rpm 1 min.
14. Discard liquid in collection tube.
15. Spin 14000 rpm 1 min one more time (dry spin).
16. Discard collection tube (but not the column).
17. (Optional: 2nd dry spin into clean collection tube.)
18. Place column in a clean labeled microtube.
19. Add 10 uL (min 6 uL for higher DNA concentration) of sterile DDH2O to top of column. Water should be pipetted directly onto center of filter.
20. Incubate at RT 1 min (or longer).
21. Spin 1 min at 14000 rpm. Discard the column.
22. Measure the concentration on the nanodrop. (You may recover the 1uL from the nanodrop if needed.)

Gel Extraction Protocol using QIAquick Gel Extraction Kit:

1. Cut the gel to separate analytical and extraction gel; place analytical gel in UV illuminator. 
2. Look at the gel under low wavelength UV (high wavelengths will denature DNA). Quickly take a polaroid image and shut OFF the UV. 
3. Cut extraction gel under white light; avoid UV illuminating the extraction gel as this drastically decreases the DNA yield. If necessary, stain with Methyl Blue. 
4. Place the cut bands in 2ml Eppendorf tubes; Weigh slices; No more than 400mg per tube
5. Add 3 volumes (6 volumes if you are afraid of getting a low yield) of Buffer QG to 1 volume of gel (100mg ~ 100ul)
6. Incubate at 50C for 10min or until gel is dissolved; vortex every 2-3 min
7. Confirm that color of mixture is yellow (if not, add 10ul of 3M NaAc, pH 5.0)
8. Add 1 gel volume of isopropanol
9. Add max of 770ul to QIAquick column and centrifuge for 1 min (max speed, ~13,000rpm, RT)
10. Run flow-through over column one more time.
11. After the second time, discard flow-through and place column back in tube.
12. If needed, add rest of mixture to same tube (up to additional 770ul), spin, and discard flow-through
13. Add 500uL of Buffer QG to column and centrifuge for 1 min (wash).
14. Wash: add 0.75ml Buffer PE (make sure that the buffer has ethanol added to it) to column.  Let stand for 2-5 minutes and then centrifuge for 1 min
15. Discard flow-through & centrifuge for 1 min
16. Place column into clean Eppendorf tube
17. Add 50ul Buffer EB or water to center of membrane.  Make sure to use warm EB (50C).  (Use 30uL if worried about low concentration.)
18. Let stand at RT for 4 min
19. Centrifuge for 1 min
20. Measure the concentration using the UV spectrophotometer.

Pro Tips

1. You don't need 2 lanes if you aren't putting your gel under UV light (the blue light and SYBR safe is fine)
2. You can up the IPA to 1/4 of the total volume
3. Warm EB (50 mL conical filled w/ water, plop the tube inside, put it in the heat block)
4. Don't let it stand at room temperature, you can do it at 5 degrees (heat block)

Gel Extraction Protocol using QIAgen MinElute Kit:

1. Cut the gel to separate analytical and extraction gel; place analytical gel in UV illuminator. 
2. Look at the gel under low wavelength UV (high wavelengths will denature DNA). Quickly take a polaroid image and shut OFF the UV. 
3. Cut extraction gel under white light; avoid UV illuminating the extraction gel as this drastically decreases the DNA yield. If necessary, stain with Methyl Blue. 
4. Place the cut bands in 2ml Eppendorf tubes; Weigh slices; No more than 300mg per tube
5. Add 3 volumes of Buffer QG to 1 volume of gel (100mg ~ 100ul)
6. Incubate at 50C for 10min or until gel is dissolved; vortex every 2-3 min
7. Confirm that color of mixture is yellow (if not, add 10ul of 3M NaAc, pH 5.0)
8. Add 1 gel volume of isopropanol
9. Add max of 800ul to MinElute column and centrifuge for 1 min (speed >= 10,000 G, RT)
10. Discard flow-through and place column back in tube.
11. If needed, add rest of mixture to same tube (up to additional 770ul), spin, and discard flow-through
12. Add 500 uL of buffer QG and spin column for 1 min and discard flow-through
13. Wash: add 0.75ml Buffer PE(make sure that the buffer has ethanol added to it) to column and centrifuge for 1 min
14. Discard flow-through & centrifuge for 1 min
15. Place column into clean Eppendorf tube
16. Add 10ul Buffer EB (10 mM TrisCl,pH 8.5) or water to center of membrane
17. Let stand at RT for 1 min
18. Centrifuge for 1 min
19. Measure the concentration using the UV spectrophotometer.

50 ng of each piece of DNA being joined

Use nanodrop to find concentration in ng/ul, then divide 50 by that concentration to find the required volume of DNA: Conc: x ng/uL Vol: 50/x uL

NOTE: If GGDonr is too concentrated, dilute it with EB or water. NOTE: Ligase buffer does not like to be freeze-thawed, so use one-time-use aliquots.

x1 uL of DNA1 x2 uL of DNA2

y uL (100ng) Donor 2ul 10X T4 Ligase Buffer 2ul 10X BSA 1ul BsaI (enzyme) HC (high concentration) 1ul T4 Ligase (enzyme) HC (high concentration)

fill to 20uL with SDIH20 (put water in before the buffer and enzymes) ------------- 20ul total

(NOTE: Make sure that Buffer and Enzyme added last, enzyme after buffer)

Take a p20, set it to 10uL and then pipet up and down.

Example: Excel File

THERMOCYCLER:

(Protocol EBGG)

37C for 5min

Part 1 50X: 37C for 2.5min 4C for 0.5min 16C for 5.5min

Part 2 37C for 10 min 80C for 20 min

4C hold (for 8+ hours)

(Check protocol by looking up the paper or other online GG protocols)

MINIPREP:

• Prepare culture in a 15 mL, round bottom tube.
• Add 5mL LB using a seriological pipette
• Add 5uL of 1000x antibiotic (Ampicillin, Kanamycin, etc.)
• Pick colony using a 10 ul pipette tip on a p2. Eject tip into tube (tip should remain in tube).
• if growing from another liquid culture, 100 uL should be plenty (replacing the 1 colony). Almost no amount is too small, just ensure that you get cells.

Appending Prefix and Suffix

Things to keep in mind!

• Annealing temperature of primers (T_m) should be around 60 C and both primers should be within 1-2C of each other
• Check the secondary structure of the primers before you order them!
  • no individual secondary structures i.e. hairpins
  • no heterostructure with the forward and reverse primers together
  • free energy of primers should be greater than -4 kCal 
  • GC content should be around 50% (40-60% is okay)

1. Dilute your DNA to the following concentrations:

1. Set up a small box (e.g. empty pipette tip box) with ice and water. Your DNA and polymerase mix will go into this box before going into the the thermocycler in order to limit endonucelase activity. 
2. Add the following DNA to a labeled 0.6ml PCR tube 

1. Program the thermocycler as follows

1. Wait for thermocycler to heat up 
2. Add 22.5uL of polymerase mix (Phusion Master Mix) to your DNA. Mix well and spin down. Transfer tubes to ice as soon as possible. 
3. Once the thermocycler has heated up to the right temperature (it should be paused at 98C), add tubes to thermocycler and resume PCR program. 

From UTexas Protocol:

Materials:

• Buffer
• Primer

You will first make a 100 uM master stock:

1. spin down primer tube in case there is some primer stuck in cap
2. Calculate amount of buffer needed- multiply nmoles of primer by 10
3. add this many uL of Buffer to dry DNA pellet
4. vortex until DNA is dissolved
5. spin down to recover drops from cap and sides of tube.

Digestion Protocol

• 20 uL Total
  • 500-1000 ng DNA (volume depends on concentration determined by nanodrop)
  • 1 uL enzyme (keep on ice, and add last!)
  • 2 uL enzyme buffer (which buffer depends on the specific enzyme)
  • fill rest with water
• Pipette up and down thoroughly to mix
• Incubate at 37 degrees for 1-3 hours
• 4-5 ul loading dye
• gel + ladder!

Tips:

Also run a non-digested construct as a control on the digestion itself.

Remember to upload gel to the wiki. Documenting your lab work is required (it is part of your lab notebook). Also remember to keep your page well organized. For ease of calculation (and for keeping track of what is where), use the digest template File:Template Digest.xlsx

From NEB:

One unit is defined as the amount of enzyme required to digest 1 µg of λ DNA in 1 hour at 37°C in a total reaction volume of 50 µl.

So check calculate how much DNA you have and use the right amount of enzyme. Or more.

This protocol is designed to purify single- or double-stranded DNA fragments from PCR and other enzymatic reactions (see page 8). For cleanup of other enzymatic reactions, follow the protocol as described for PCR samples or use the MinElute Reaction Cleanup Kit. Fragments ranging from 100 bp to 10 kb are purified from primers, nucleotides, polymerases, and salts using QIAquick spin columns in a microcentrifuge.

Important points before starting

• Add ethanol (96–100%) to Buffer PE before use (see bottle label for volume).
• All centrifugation steps are carried out at 17,900 x g (13,000 rpm) in a conventional tabletop microcentrifuge at room temperature.
• Add 1:250 volume pH indicator I to Buffer PB (i.e., add 120 µl pH indicator I to 30 ml Buffer PB or add 600 µl pH indicator I to 150 ml Buffer PB). The yellow color of Buffer PB with pH indicator I indicates a pH of ≤ 7.5.
• Add pH indicator I to entire buffer contents. Do not add pH indicator I to buffer aliquots.
• If the purified PCR product is to be used in sensitive microarray applications, it may be beneficial to use Buffer PB without the addition of pH indicator I.

Procedure

1. Add 5 volumes of Buffer PB to 1 volume of the PCR sample and mix. It is not necessary to remove mineral oil or kerosene.

   For example, add 500 µl of Buffer PB to 100 µl PCR sample (not including oil).

2. If pH indicator I has beein added to Buffer PB, check that the color of the mixture is yellow.

   If the color of the mixture is orange or violet, add 10 µl of 3 M sodium acetate, pH 5.0, and mix. The color of the mixture will turn to yellow.

3. Place a QIAquick spin column in a provided 2 ml collection tube.
4. To bind DNA, apply the sample to the QIAquick column and centrifuge for 30–60 s.

5. Discard flow-through. Place the QIAquick column back into the same tube.

   Collection tubes are re-used to reduce plastic waste.

6. To wash, add 0.75 ml Buffer PE to the QIAquick column and centrifuge for 30–60 s.

7. Discard flow-through and place the QIAquick column back in the same tube. Centrifuge the column for an additional 1 min.

   IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is discarded before this additional centrifugation.

8. Place QIAquick column in a clean 1.5 ml microcentrifuge tube.

9. To elute DNA, add 50 µl Buffer EB (10 mM Tris·Cl, pH 8.5) or water (pH 7.0–8.5) to the center of the QIAquick membrane and centrifuge the column for 1 min. Alternatively, for increased DNA concentration, add 30 µl elution buffer to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge.

   IMPORTANT: Ensure that the elution buffer is dispensed directly onto the QIAquick membrane for complete elution of bound DNA. The average eluate volume is 48 µl from 50 µl elution buffer volume, and 28 µl from 30 µl elution buffer.

   Elution efficiency is dependent on pH. The maximum elution efficiency is achieved between pH 7.0 and 8.5. When using water, make sure that the pH value is within this range, and store DNA at –20°C as DNA may degrade in the absence of a buffering agent. The purified DNA can also be eluted in TE buffer (10 mM Tris·Cl, 1 mM EDTA, pH 8.0), but the EDTA may inhibit subsequent enzymatic reactions.

10. If the purified DNA is to be analyzed on a gel, add 1 volume of Loading Dye to 5 volumes of purified DNA. Mix the solution by pipetting up and down before loading the gel.

    Loading dye contains 3 marker dyes (bromophenol blue, xylene cyanol, and orange G) that facilitate estimation of DNA migration distance and optimization of agarose gel run time. Refer to Table 2 (page 15) to identify the dyes according to migration distance and agarose gel percentage and type.

For PCR Primers in Geneious:

• Highlight the gene that you want to amplify
• Select "Primers" from the top banner and "design new primers" from the drop down tab
• Make sure you have checked "Forward Primer" and "Reverse Primer"
• Set task to "precise" if you are doing this for a cloning reaction
• check "Target Region"
• Modify characteristics as needed - see checking Tm and other primer design tips in the protocols page
• Hit OK and pick the best primer generated - hovering your mouse over the primer will give you Tm and GC content
• Check your PCR reaction coditions, Geneious may not have an accurate model for predicting Tm, check the NEB website.

For Sequencing Primers:

• You will get some amount of good read from a primer, look this up. Some primers are standard, they normally read from the ends of the insert. Check the backbone to see what sequencing primers are in place. These may be available from the sequencing shop, or they should at least be in the fridge.
• If your gene is less than whatever length you can sequence it using back bone primers and do not need to design your own
• If your gene is larger then what can be read by the back bone primers, you will need to sequence part way through with custom primers.
• Highlight a 50-100 bp region that you want the primer to bind in
• Select "Primers" from the top banner and "design new primers" from the drop down tab
• Set task to "Generic"
• Check forward primer and DO NOT check reverse primer
• Check "Included Region"
• Set "Tm Min" to 50 optimal to 55 and "Tm Max" to 60
• Set "GC Min" to 45 optimal to 50 and "GC Max to 55
• Set "GC Clamp" to 1
• Hit OK and pick the best primer generated - hovering your mouse over the primer will give you Tm and GC content

General tips for sequencing:

Reads are only clear about 30 bp after the primer to 600 bp after the primer.

The easiest way to pick primers is to start with one in a 50 bp region in front of the gene and then measure out 500 bp from that primer then place the next one in the following 100 bp region then measure out 500 bp and place one in the next 100 bp region and so on.

If you are planning on doing additional cloning with this part try to make some of the primers usable for future constructs

For Both:

• Once your primers are designed. right click on the primer annotation and select "Extract Regions" this will make a primer file in the same folder
• Select all of your primers and then click File => Export
• Export the documents as a .csv
• Highlight only name and sequence
• Email the csv to your supervisor to be ordered

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 35.

Note: All protocol steps should be carried out at room temperature.

Procedure

1. Resuspend pelleted bacterial cells in 250 µl Buffer P1 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.

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.

8. Wash QIAprep Spin Column by adding 0.75 ml Buffer PE and centrifuging for 30–60 s.

9. Discard the flow-through, and centrifuge for an additional 1 min to remove residual wash buffer.

   IMPORTANTResidual 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

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.

The QIAprep Spin Miniprep procedure can be performed using 5 ml centrifuge tubes (e.g., Greiner, Cat. No. 115101 or 115261) as collection tubes to decrease handling.

Note: All protocol steps should be carried out at room temperature.

Procedure

1. Resuspend pelleted bacterial cells in 250 µl Buffer P1 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. Place a QIAprep Spin Column in a 5 ml centrifuge tube instead of a 2 ml collection tube.

5. Apply the supernatants from step 4 to the QIAprep Spin Column by decanting or pipetting.

6. Centrifuge at 3000 x g for 1 min using a suitable rotor (e.g., Beckman® GS-6KR centrifuge at ~4000 rpm). (The flow-through does not need to be discarded.)

7. (Optional): Wash the QIAprep Spin Column by adding 0.5 ml Buffer PB and centrifuging for 30–60 s. Discard the flow-through.

   For washing steps, centrifugation should be performed at 3000 x g for 1 min. The flow-through does not need to be discarded.)

8. Wash QIAprep Spin Column by adding 0.75 ml Buffer PE and centrifuging for 30–60 s.

   For washing steps, centrifugation should be performed at 3000 x g for 1 min. The flow-through does not need to be discarded.)

9. Transfer the QIAprep Spin Column to a microcentrifuge tube. Centrifuge at maximum speed for 1 min. Continue with step 10 of the protocol.

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.