Manual / Product Insert

User Guide: TURBO DNA-free Kit (English )

Version: H
Catalog #
  • AM1907
  • AM1907M(Discontinued)

Product FAQ

How do I remove bubbles from the vials of either ProLong Antifade Mountant, ProLong Gold, ProLong Diamond or ProLong Glass Antifade Mountants or ProLong-mounted samples?

Answer

Bubbles may be removed by one of two methods:

1. Place the amount of ProLong reagent you wish to use on your sample (plus a little excess) in a microcentrifuge tube. Close the cap and centrifuge this aliquot using a tabletop microcentrifuge (speed from 7, 000 to 13,000 rpm). Bubbles should move to the top and these bubbles may be aspirated using a pipettor/pipette tip.
2. Unscrew the lid of the bottle/vial containing the ProLong reagent to make it loose, but do not remove the lid. Place the entire bottle/vial into a vacuum flask, using a faucet aspirator (faucet T-tube). Apply a vacuum (water running through the faucet) and allow vacuum aspiration to occur from 10 to 20 minutes to degas the mixture.

To avoid the formation of bubbles on a sample or to remove bubbles:

1. Before pipetting the desired amount of ProLong reagent for mounting, set the pipettor for a slight excess volume. When pipetting up the mixture, do not pipette up the complete amount, but lift up the pipette tip from the bottle with the pipettor not yet up to full volume. This prevents the aspiration of bubbles into the pipette tip.
2. Bubbles trapped during application of the coverslip: When placing your coverslip onto your drop of ProLong reagent, place the coverslip at a slight angle then, gently lower the coverslip. If the coverslip is lowered flat onto the sample, or lowered too quickly, bubbles can be trapped.
3. Bubbles trapped in tissue: One problem with tissue sections, particularly cryosections is that air can get trapped within and under the section. Upon mounting, bubbles are not observed but as the mountant hardens, it compresses the sample slightly, forcing air out of the section. This leads to microscopic bubbles forming over the section, trapped within the mountant. To avoid this, degas the tissue sample prior to mounting. Place the sections submerged in buffer or blocking solution, into a vacuum chamber and expose the sample to the vacuum. This will degas the sections and buffer. Remove the sample from this degassed buffer and mount.
4. If ProLong-mounted samples have already cured but have bubbles, you can un-mount your sample by placing the slides into PBS (Coplin jar or a Petri dish filled with PBS). The ProLong reagent will swell and the coverslip will slide off or can be gently removed manually. You can then re-mount with a new aliquot of ProLong reagent.

Answer Id: E14794

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Product FAQ

What is the smallest sample volume I can use when extracting RNA with TRIzol Reagent?

Answer

Small volumes (0.5-0.8 mL) of TRIzol Reagent have been used successfully for 10^2 to 10^5 cells, but if small volumes are used, we recommend using smaller tubes in order to have the tallest possible column of aqueous phase. The taller the column of liquid, the less likely that contamination from the interphase will occur.

Here is a protocol for isolation of RNA from small quantities of tissue (1-10 mg) or cells (100-10,000):
1. Add 800 µL TRIzol Reagent to the sample. Homogenize cells by pipetting repeatedly. Add 200 µg glycogen (Cat. No. 10814010) directly to the TRIzol Reagent. If processing tissue, pulverize in liquid nitrogen first and then add 800 µL TRIzol Reagent containing 200 µg glycogen (final concentration 250 µg/mL) followed by vigorous vortexing or power homogenization.
2. Place at room temperature, cap the vial, and vortex at high speed for 10 seconds. Make sure the TRIzol Reagent wets the side of the vial in order to solubilize any sample that may be remaining on the walls.
3. Shear the genomic DNA in the sample by passing twice through a 26-gauge needle connected to a 1 mL syringe. Using the syringe, transfer the sample to a sterile 1.5 mL microcentrifuge tube.
4. Add 160 µL of chloroform (or 49:1 chloroform:isoamyl alcohol) to each sample and vortex up to 30 seconds. Centrifuge at maximum speed in a microcentrifuge for 5 minutes to separate the phases.
5. Transfer the upper aqueous phase to a fresh tube and add 400 µL ice-cold isopropanol. Allow the samples to precipitate at -20 degrees C for 1 hour to overnight. Pellet the RNA by centrifugation at maximum speed in the microfuge for 15 minutes at room temperature.
6. Decant the supernatant. Wash the pellet in 200 µL of 70% ethanol and centrifuge again for 10 minutes at maximum speed. Decant the supernatant, removing as much as possible without disturbing the pellet. Dry the RNA pellet.
7. Resolubilize the pellet in 30-50 µL RNAse-free deionized water. If tissue is high in RNAses (e.g., adrenal gland, pancreas), resuspend in 100% deionized formamide. Be sure to vortex or pipette the sample up and down to ensure that the pellet is fully resolubilized. Store at -70 degrees C.

Answer Id: E7833

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Product FAQ

How do I process plasma samples for use on the Luminex assay platform?

Answer

Separate the cells from the plasma samples by centrifugation at 2,000 x g for 10 min in a refrigerated centrifuge. Centrifugation at this force is necessary to deplete the sample of platelets. Transfer the supernatant to a chilled clean polypropylene tube with a sterile Pasteur pipette. Maintain the samples at 2-8 degrees C while handling.

If the plasma is to be analyzed at a later date, apportion it into aliquots in polypropylene microcentrifuge tubes and store at -80 degrees C. Avoid multiple freeze-thaw cycles. When you are ready to analyze them, allow the samples to thaw on ice. All plasma samples should be clarified by centrifugation (14,000 rpm for 10 min at 4 degrees C) in a refrigerated microcentrifuge immediately prior to analysis. Follow the assay procedure provided with the kit for appropriate dilutions.

Answer Id: E12641

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Product FAQ

What is the recommended protocol for preparation of competent yeast for large-scale transformation with bait and plasmid-library in the ProQuest system?

Answer

(1) Suspend several colonies of MaV203 in 50 µL autoclaved, distilled water in a microcentrifuge tube and spread it onto the center of a 10-cm YPAD plate using an autoclaved loop or toothpick. Repeat procedure for a second YPAD plate. Incubate both plates for 18-24 h at 30 degrees C.

(2) Scrape and completely suspend the cells (by brief vortexing and pipetting up and down) in 10 mL autoclaved, distilled water. Add a sufficient volume of this cell suspension to two 1-L flasks each containing 500 mL liquid YPAD medium to give an OD600 of ~0.1. Reserve approximately 10 mL YPAD medium to use as a blank in the spectrophotometer.
Note: Perform serial 1:10 dilutions in water of the 10-mL cell suspension then determine the OD600 of each dilution to allow an estimate of cell suspension required to produce the desired OD of 0.1. Appropriate cell densities require that the measured OD be <1.0. Verify that the OD is ~0.1 after inoculation. Use plastic cuvettes for all OD600 measurements.

(3) Shake (~250 rpm) at 30 degrees C until the OD600 reaches ~0.4 (usually ~5 h). Read the OD.

(4) Prepare fresh:
225 mL 1X TE/LiAc by combining 22.5 mL 10X TE, 22.5 10X LiAc, and 180 mL autoclaved H2O.
30 mL PEG/LiAc by combining 3 mL 10X TE, 3 mL 10X LiAc, and 24 mL 50% PEG-3350.
200 µL carrier DNA by boiling sonicated herring sperm DNA or sonicated salmon sperm DNA (10 mg/mL) for 5 min and placing on ice until use.

(5) Split each 500 mL of yeast cells into two conical 250-mL tubes and centrifuge at 3,000 x g for 5 min at room temperature.

(6) Pour off the supernatants and gently suspend each pellet by pipetting up and down in 100 mL autoclaved, distilled water at room temperature.

(7) Centrifuge at 3,000 x g for 5 min at room temperature. Pour off the supernatant of the centrifuged cells and suspend each cell pellet in 50 mL 1X TE/LiAc solution.

(8) Centrifuge at 3,000 x g for 5 min at room temperature. Carefully pour off the supernatants and suspend each cell pellet in a final volume of 1 mL 1X TE/LiAc solution and pool all suspensions for a total of 4 mL.

(9) Perform 30 transformations. Combine 4 mL of cells, 200 µL freshly boiled carrier DNA and 150 µg (~1 µg/µL) bait plasmid DNA and 150 µg (~1 µg/µL) plasmid-library plasmid DNA. Mix gently by pipetting up and down. Add 24 mL PEG/LiAc solution and mix gently, but completely. Aliquot into 30 autoclaved microcentrifuge tubes of 950 µL each.

(10) Incubate for 30 min in a 30 degrees C water bath.

(11) Heat shock for 15 min in a 42 degrees C water bath.

(12) Centrifuge in a microcentrifuge (6,000 - 8,000 x g) for 20-30 s at room temperature. Carefully remove the supernatant.

(13) Gently suspend each pellet in 400 µL autoclaved, distilled water by pipetting up and down.

(14) To estimate the total number of transformants, plate two dilutions of the transformation. Mix 10 µL of transformation with 90 µL autoclaved, distilled water. Plate 100 µL on a 10-cm SC-Leu-Trp plate (1:800 final dilution factor). Mix 10 µL of transformation with 990 µL autoclaved, distilled water. Plate 100 µL on a 10-cm SC-Leu Trp plate (1:8,000 final dilution factor).

Answer Id: EF

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Product FAQ

How do I process cell culture supernatants for use on the Luminex assay platform?

Answer

Cells should be in log-phase growth. Stimulate cells as desired in appropriate cell culture flasks. Using sterile technique, remove the desired volume of conditioned cell culture medium with a pipette and transfer the medium to clean polypropylene microcentrifuge tubes. Centrifuge the medium at 14,000 rpm for 10 min at 4 degrees C in a refrigerated microcentrifuge to remove any cells or cellular debris. Aliquot the clarified medium into clean polypropylene microcentrifuge tubes. These samples are ready for the assay. Alternatively, clarified medium samples can be aliquoted and stored at -80 degrees C for future analysis. Avoid multiple freeze-thaw cycles. Frozen samples should be allowed to thaw on ice just prior to running the assay. Thawed samples should be clarified by centrifuging at 14,000 rpm for 10 min at 4 degrees C in a refrigerated microcentrifuge prior to analysis to prevent clogging of the Luminex probe and/or filter plate. Follow the assay procedure provided with the kit for appropriate dilutions.

Answer Id: E12642

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Product FAQ

How can I isolate DNA from Thermo Fisher Scientific TBE and TBE-Urea gels?

Answer

Here is the protocol you can follow:
(1) Run the gel. Locate the DNA of interest by autoradiography or by examination of ethidium bromide stained gels.
(2) Use a razor blade to cut out the upper 1/2 to 2/3 of the band of interest.
To recover a fragment of DNA identified by autoradiography, cut out from the X-ray film a small rectangle encompassing the image of the fragment. Align the film over the gel and cut out the segment of polyacrylamide bordered the rectangular hole in the film.
(3) Transfer the gel slice to a microcentrifuge tube. Use a sterile glass rod to crush the slice against the wall of the tube.
(4) Calculate the volume of the slice and add 1 to 2 volumes of elution buffer to the microcentrifuge tube.
Elution buffer: 0.5 M ammonium acetate, 10 mM magnesium sulfate, 1 mM EDTA (pH 8.0), 0.1% SDS
It is convenient if the volume of elution buffer is no greater than 0.5 mL since the eluted fragment of DNA can then be precipitated with ethanol in a single tube.
(5) Close the tube and incubate at 37 degrees C on a rotary platform. Small fragments of DNA (500 bases) are eluted in 3 to 4 hours.
(6) Centrifuge the sample at 12,000 g for 1 minute at 4 degrees C in a microcentrifuge. Transfer the supernatant to a fresh microcentrifuge tube, being careful to avoid transferring fragments of polyacrylamide.
(7) Add additional 0.5 mL volume of elution buffer to the pellet of polyacrylamide, vortex briefly, and re-centrifuge. Combine the two supernatants.
(8) Remove any remaining fragments of polyacrylamide by passing the supernatant through a disposable plastic column or a syringe barrel containing Whatman GF/C filter or packed siliconized glass wool.
(9) Add 2 volumes of ethanol at 4 degrees C and store the solution on the ice for 30 min. Recover the DNA by centrifugation at 12,000 g for 10 min at 4 degrees C in a microcentrifuge.
(10) Re-dissolve the DNA in 200 µL of Tris-EDTA (TE) buffer (pH 7.6), add 25 µL of 3M sodium acetate (pH 5.2) and re-precipitate the DNA with 2 volumes of ethanol as described in step 9. TE Buffer (pH 7.6): 10 mM Tris HCl (pH 7.6), 1mM EDTA (pH 8.0).
(11) Carefully rinse the pellet once with 70% ethanol and redissolve the DNA in TE buffer (pH 7.6) to a final volume of 10 µL.
(12) Check the amount and the quality of the fragment by polyacrylamide gel electrophoresis.

Answer Id: E4566

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Product FAQ

How do you recommend that I prepare my DNA for successful electroporation of E. coli?

Answer

For best results, DNA used in electroporation must have a very low ionic strength and a high resistance. A high-salt DNA sample may be purified by either ethanol precipitation or dialysis.

The following suggested protocols are for ligation reactions of 20ul. The volumes may be adjusted to suit the amount being prepared.

Purifying DNA by Precipitation: Add 5 to 10 ug of tRNA to a 20ul ligation reaction. Adjust the solution to 2.5 M in ammonium acetate using a 7.5 M ammonium acetate stock solution. Mix well. Add two volumes of 100 % ethanol. Centrifuge at 12,000 x g for 15 min at 4C. Remove the supernatant with a micropipet. Wash the pellet with 60ul of 70% ethanol. Centrifuge at 12,000 x g for 15 min at room temperature. Remove the supernatant with a micropipet. Air dry the pellet. Resuspend the DNA in 0.5X TE buffer [5 mM Tris-HCl, 0.5 mM EDTA (pH 7.5)] to a concentration of 10 ng/ul of DNA. Use 1 ul per transformation of 20 ul of cell suspension.

Purifying DNA by Microdialysis: Float a Millipore filter, type VS 0.025 um, on a pool of 0.5X TE buffer (or 10% glycerol) in a small plastic container. Place 20ul of the DNA solution as a drop on top of the filter. Incubate at room temperature for several hours. Withdraw the DNA drop from the filter and place it in a polypropylene microcentrifuge tube. Use 1ul of this DNA for each electrotransformation reaction.

Answer Id: E4159

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Product FAQ

How do I process cell lysates (extract cellular proteins) for the Luminex assay platform?

Answer

The protocols mentioned below have been applied to several human and mouse cell lines. You should optimize the cell extraction procedures for your own applications.

Cell Lysis Procedure

Non-adherent cells:
Pellet cells by low-speed centrifugation. Remove the medium from the pellet, and wash twice with ice-cold PBS. Remove the PBS, and resuspend the cell pellet in cell lysis buffer (recommended cell lysate concentration is 2-5 mg/mL). Incubate 15 min on ice with occasional vortexing. Transfer the lysate to a microcentrifuge tube and centrifuge at 14,000 rpm for 10 min at 2-8 degrees C. Aliquot the cleared lysate into clean microcentrifuge tubes, and determine total protein concentration.

Adherent cells:
Remove the tissue culture medium from the cells, and wash twice with ice-cold PBS. Remove the PBS, add cell lysis buffer (recommended cell lysate concentration is 2-5 µg/mL), and incubate 15 min on ice. Collect the cell lysate and transfer it to a microcentrifuge tube. Centrifuge at 14,000 rpm for 10 min at 2-8 degrees C. Aliquot the cleared lysate into clean microcentrifuge tubes, and determine total protein concentration. Lysates should be frozen and stored at -80 degrees C or analyzed shortly after collection. Avoid multiple freeze-thaw cycles of frozen samples. Thaw completely, mix well, and clarify by centrifugation (14,000 rpm for 5 min) prior to analysis to prevent clogging of the filter plates.

Recommended Cell Lysis Buffer:
NP40 Cell Lysis Buffer (Cat. No. FNN0021) Note: Lysates prepared with NP40 Lysis Buffer must be diluted at least 5-fold prior to running the assay. The recipe to make the buffer is as follows: 50 mM Tris, pH 7.4, 50 mM NaF, 260 mM NaCl, 1 mM Na3VO4, 5 mM EDTA, 0.02% NaN3, 1% Nonidet P40. The NP40 Cell Lysis Buffer (without protease inhibitor cocktail and PMSF) is stable for 2-3 weeks at 2-8 degrees C, or for 6 months when stored in aliquots at -20 degrees C.

Add fresh to the NP40 Cell Lysis Buffer just before use:
1 mM PMSF (stock 0.3 M in DMSO)
Protease inhibitor cocktail (Sigma, Cat. No. P-2714)
Alternate Cell Extraction Buffer
Cell Extraction Buffer (Cat. No. FNN0011) Note: Lysates prepared with Cell Extraction Buffer must be diluted at least 10-fold prior to running the assay. Or 10 mM Tris, pH 7.4 2 mM Na3VO4 100 mM NaCl 1% Triton X-100 1 mM EDTA 10% glycerol 1 mM EGTA 0.1% SDS 1 mM NaF 0.5% deoxycholate 20 mM Na4P2O7 The Cell Extraction Buffer (without protease inhibitor cocktail and PMSF) is stable for 2-3 weeks at 2-8 degrees C, or for 6 months when stored in aliquots at -20 degrees C. Add fresh to the Cell Extraction Buffer just before use: 1 mM PMSF (stock 0.3 M in DMSO) Protease inhibitor cocktail (Sigma, Cat. No. P-2714)

Answer Id: E12651

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Product FAQ

What is your recommended protocol for generating ss phagemid DNA?

Answer

The following protocol can be used to prepare ssDNA from DH12S or DH5aF', DH5aF'IQ, DH11S cells (this strain not currently offered for sale). Use a stock of the helper phage M13KO7 (1) that is of known titer. For convenience, DH12S cells are supplied with M13KO7 helper phage. DH12S cells are both endA positive, so single stranded DNA isolated from these strains tend to be less contaminated with double stranded rf DNA.

Small-Scale Preparation of ss Phagemid DNA:
1. Pick a single colony of cells containing phagemid DNA and resuspend in 2 mL TBG (1.2% tryptone, 2.4% yeast extract 0.4% glycerol, 17 mM KH2PO4 and 55 mM KH2PO4 and 20 mM glucose) containing 100 µg/mL ampicillin in a 15 mL tube.
2. Immediately add 10 µL M13KO7 helper phage stock at 10E11 pfu/mL.
3. Incubate cells at 37 degrees C with vigorous agitation (275 rpm) for 2 hrs.
4. Add kanamycin to a final concentration of 75 µg/mL and incubate cells at 37 degrees C with vigorous agitation (275 rpm) for 18 to 24 hrs.
5. Transfer 1.5 mL of culture to a sterile microcentrifuge tube and pellet cells by centrifuging at 14,000 x g for 10 min at 4 degrees C.
6. Transfer supernatant to fresh microcentrifuge tube and repeat the centrifugation.
7. Transfer 1.2 mL supernatant to a fresh microcentrifuge tube and add 300 µL of 2.5 M NaCl in 40% PEG 4000.
8. Vortex and incubate on ice for 15 min.
9. Centrifuge at 14,000 x g for 15 min at 4 degrees C.
10. Resuspend the pellet in 50 µL TE and phenol extract to remove the viral coat.
11. Use 10 µL of the final 50 µL volume for gel analysis.
This protocol yields 0.5 to 1 µg ss phagemid DNA.

Large-Scale Preparation of ss Phagemid DNA:
1. Resuspend a single colony in 5 mL of TB or LB broth containing 100 µg/mL ampicillin in a 15-mL tube.
2. Shake at 37 degrees C and 275 rpm overnight.
3. Add 100 µL of the overnight culture to 200 mL LB broth and 100 µg/mL ampicillin in a 1 L flask. Incubate at 37 degrees C with shaking (275 rpm) for 3 hrs.
4. Add 200 µL of M13KO7 helper phage (1 x 10E11 pfu/mL) to the culture and continue to incubate for 2 hrs.
5. Add 1.5 mL of 1% (w/v) kanamycin to the cells for a final concentration of 75 µg/mL. Incubate the infected cells for an additional 18 to 24 h at 37 degrees C.
6. Centrifuge this culture at 16,000 x g for 15 min at 4 degrees C .
7. Filter the supernatant through a 0.2 µm sterile filter into an autoclaved centrifuge bottle. Add 40 µL of DNase I (50 units/µL) and incubate at room temperature for 3 hrs. This step should remove any residual ds plasmid DNA contamination.
8. Transfer 100 mL of the supernatant to another centrifuge bottle. Add 25 mL of 2.5 M NaCl in 40% PEG 4000 to each of the centrifuge bottles containing the supernatant.
9. Vortex the mixture, incubate on ice for 1 h, and centrifuge at 16,000 x g for 20 min at 4 degrees C.
10. Carefully discard the supernatant. To fully drain off the remaining solution from the pellets, place the bottles on an angle, with the pellet side facing up for 10 to 15 min. Remove the solution with a sterile Pasteur pipet.
11. Resuspend the pellets in 2 mL of TE buffer. Add 10 µL of proteinase K solution (20 mg/mL), 20 µL of 10% SDS, and incubate this mixture at 45 degrees C for 1 hr.
12. Transfer the digested mixture to three microcentrifuge tubes and extract four times with an equal volume of phenol:chloroform: isoamyl alcohol (25:24:1), precipitate with ethanol, and dissolve in 100 µL TE buffer.
13. Freeze the solubilized DNA at -20 degrees C for 1 hr and centrifuge in a microcentrifuge at 14,000 x g for 15 min at 4 degrees C.
14. Transfer the supernatant containing the ss plasmid DNA to a fresh tube and discard the polysaccharide pellet. Store the ssDNA at 4 degrees C.
15. Determine the DNA concentration (OD260).
This protocol yields ~100 to 200 µg ss phagemid DNA.

(1) Vieira, J. and Messing, J. (1987) Methods in Enzymology 153, 3.

Answer Id: E4161

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Product FAQ

How can I extract peptides from NuPAGE gels for MALDI/MS analysis?

Answer

To extract peptides from NuPAGE gels perform the following protocol:

1) Dehydrate the gel band in 100% methanol for 5 min at room temperature.

2) Rehydrate the gel band in 30% methanol for 5 min.

3) Wash the gel band twice in ultrapure water for 10 min.

4) Wash the gel band three times with 100 mM ammonium bicarbonate containing 30% acetonitrile for 10 min.

5) After the last wash, cut the gel into small pieces and wash the gel pieces in ultrapure water.

6) Dry the gel pieces in a SpeedVac concentrator for 30 min

7) Resuspend the gel pieces in 50 mM ammonium bicarbonate. Add approximately 5 µL buffer per square millimeter of gel. There should be sufficient buffer to cover the gel pieces.

8) Add 5-10 ng/µL trypsin and incubate overnight at 37°C.

9) Centrifuge at maximum speed in a microcentrifuge for 1 min and transfer supernatant to sterile microcentrifuge tube using a clean pipette tip.

10) Extract remaining peptides from the gel with 10-20 µL 50 acetonitrile containing 0.1 trifluoroacetic acid at room temperature. Combine this extract with the supernatant from step 9.

11) Concentrate the sample from step 10 to 4-5 µL using a SpeedVac concentrator and proceed to MALDI/MS analysis. Be sure to include a control sample for MALDI/MS analysis.

Answer Id: E4145

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Product FAQ

Do you have a protocol for purification of His-tagged proteins synthesized in the ExpressWay Cell-Free Expression System?

Answer

Purification may be performed at 4 degrees C or room-temperature depending upon the sensitivity of the synthesized product.

1. Upon completion of incubation, remove the desired portion of reaction for His-tag purification to a clean microcentrifuge tube. Add 4 volumes of Binding buffer and vortex briefly (Add 200 µL for 50 µL of reaction). Centrifuge 5 minutes at 12,000 rpm.
2. Transfer the supernatant to a 2.0 mL tube containing 50 µL pre-equilibrated resin.
3. Incubate with shaking or mixing for 30-60 minutes.
4. Spin down resin for 2 minutes at 800 x g. Do not spin any higher or the resin will collapse and recovery will be low. Carefully remove supernatant.
5. Add 200 µL wash buffer and mix for 5 minutes.
6. Spin down resin for 2 minutes at 800 x g. Carefully remove supernatant.
7. Repeat steps 5 and 6.
8. Add 100 µL Elution Buffer and mix for 5 minutes.
9. Spin down resin for 2 minutes at 800 x g. Carefully remove and save supernatant.
10. Repeat steps 8 and 9.

Binding Buffer:
50 mM NaP04, pH 7.0
500 mM NaCl
6 M guanidine HCl (optional)**

Wash Buffer:
50 mM NaP04, pH 7.0
500 mM NaCl
15-25 mM imidazole*

Elution Buffer:
50 mM NaP04, pH 7.0
500 mM NaCl
150-250 mM imidazole*

**Depending on downstream applications, the purification may be performed under semi-denaturing conditions, or native conditions. Under semi-denaturing conditions, dilute the reaction in denaturing Binding Buffer containing 6 M guanidine HCl; then wash and elute with native buffers.

The concentration of imidazole is dependent upon the type of resin used. For Ni-NTA or ProBond resins, use 25 mM imidazole in the wash buffer and 250 mM imidazole in the elution buffer.

Answer Id: E12951

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Product FAQ

How do I anneal my single-stranded DNA oligos to create a ds oligo?

Answer

You will want to anneal equal amounts of the top- and bottom-strand oligos to generate the ds oligos. If your single-stranded oligos are supplied lyophilized, resuspend them in water or TE buffer to a final concentration of 200 µM before use. We generally perform the annealing reaction at a final single-stranded oligo concentration of 50 µM. Annealing at concentrations lower than 50 µM can significantly reduce the efficiency. Note that the annealing step is not 100% efficient; approximately half of the single-stranded oligos remain unannealed even at a concentration of 50 µM. Please see the steps below:

1. In a 0.5 mL sterile microcentrifuge tube, set up the following annealing reaction at room temperature.
“Top-strand” DNA oligo (200 µM) - 5 µL, “Bottom-strand” DNA oligo (200 µM)- 5 µL, 10X Oligo Annealing Buffer - 2 µL, DNase/RNase-Free Water - 8 µL which should make a total volume of 20 µL.
2. If reannealing the lacZ ds control oligo, centrifuge its tube briefly (approximately 5 seconds), then transfer the contents to a separate 0.5 mL sterile microcentrifuge tube.
3. Incubate the reaction at 95 degrees C for 4 minutes.
4. Remove the tube containing the annealing reaction from the water bath or the heat block, and set it on your laboratory bench.
5. Allow the reaction mixture to cool to room temperature for 5-10 minutes. The single-stranded oligos will anneal during this time.
6. Place the sample in a microcentrifuge and centrifuge briefly (approximately 5 seconds). Mix gently.
7. Remove 1 µL of the annealing mixture and dilute the ds oligo as directed.
8. Store the remainder of the 50 µM ds oligo mixture at -20 degrees C.
You can verify the integrity of your annealed ds oligo by agarose gel electrophoresis, if desired.

Answer Id: E9981

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