pcDNA™3.3-TOPO™ TA Cloning™ Kit - FAQs

Can I store my competent E. coli in liquid nitrogen?

We do not recommend storing competent E. coli strains in liquid nitrogen as the extreme temperature can be harmful to the cells. Also, the plastic storage vials are not intended to withstand the extreme temperature and may crack or break.

How should I store my competent E. coli?

We recommend storing our competent E. coli strains at -80°C. Storage at warmer temperatures, even for a brief period of time, will significantly decrease transformation efficiency.

What is the difference between pcDNA 3.4-TOPO TA and pcDNA 3.3-TOPO TA vectors?

pcDNA 3.4-TOPO TA vector is an improvement over pcDNA 3.3-TOPO TA vector. It contains the WPRE (Woodchuck Posttranscriptional Regulatory Element) that allows for 2- to 3-fold higher levels of expression than pcDNA 3.3-TOPO TA vector.

I performed stable selection but my antibiotic-resistant clones do not express my gene of interest. What could have gone wrong?

Here are possible causes and solutions:

Detection method may not be appropriate or sensitive enough:
- We recommend optimizing the detection protocol or finding more sensitive methods. If the protein is being detected by Coomassie/silver staining, we recommend doing a western blot for increased sensitivity. The presence of endogenous proteins in the lysate may obscure the protein of interest in a Coomassie/silver stain. If available, we recommend using a positive control for the western blot.
- Insufficient number of clones screened: Screen at least 20 clones.
- Inappropriate antibiotic concentration used for stable selection: Make sure the antibiotic kill curve was performed correctly. Since the potency of a given antibiotic depends upon cell type, serum, medium, and culture technique, the dose must be determined each time a stable selection is performed. Even the stable cell lines we offer may be more or less sensitive to the dose we recommend if the medium or serum is significantly different.
- Expression of gene product (even low level) may not be compatible with growth of the cell line: Use an inducible expression system.
- Negative clones may result from preferential linearization at a vector site critical for expression of the gene of interest: Linearize the vector at a site that is not critical for expression, such as within the bacterial resistance marker.

I used a mammalian expression vector but do not get any expression of my protein. Can you help me troubleshoot?

Here are possible causes and solutions:

- Try the control expression that is included in the kit
Possible detection problem:

- Detection of expressed protein may not be possible in a transient transfection, since the transfection efficiency may be too low for detection by methods that assess the entire transfected population. We recommend optimizing the transfection efficiency, doing stable selection, or using methods that permit examination of individual cells. You can also increase the level of expression by changing the promoter or cell type.
- Expression within the cell may be too low for the chosen detection method. We recommend optimizing the detection protocol or finding more sensitive methods. If the protein is being detected by Coomassie/silver staining, we recommend doing a western blot for increased sensitivity. The presence of endogenous proteins in the lysate may obscure the protein of interest in a Coomassie/silver stain. If available, we recommend using a positive control for the western blot. Protein might be degraded or truncated: Check on a Northern. Possible time-course issue: Since the expression of a protein over time will depend upon the nature of the protein, we always recommend doing a time course for expression. A pilot time-course assay will help to determine the optimal window for expression. Possible cloning issues: Verify clones by restriction digestion and/or sequencing.

Find additional tips, troubleshooting help, and resources within our Protein Expression Support Center.

I am using a mammalian expression vector that has the neomycin resistance gene. Can I use neomycin for stable selection in mammalian cells?

No; neomycin is toxic to mammalian cells. We recommend using Geneticin (a.k.a. G418 Sulfate), as it is a less toxic and very effective alternative for selection in mammalian cells.

Is it okay if my construct has an ATG that is upstream of the ATG in my gene of interest? Will it interfere with translation of my gene?

Translation initiation will occur at the first ATG encountered by the ribosome, although in the absence of a Kozak sequence, initiation will be relatively weak. Any insert downstream would express a fusion protein if it is in frame with this initial ATG, but levels of expressed protein are predicted to be low if there is a non-Kozak consensus sequence. If the vector contains a non-Kozak consensus ATG, we recommend that you clone your gene upstream of that ATG and include a Kozak sequence for optimal expression.

What is the difference between pcDNA3.1 vectors and the pcDNA3.3-TOPO vector?

pcDNA3.1 vectors contain the core CMV promoter that is truncated before the start of transcription, whereas the pcDNA 3.3-TOPO vector has the 672 bp native CMV promoter. This native CMV promoter allows high-level gene expression with two- to five-fold higher protein yields compared to other expression vectors. pcDNA3.1 vectors are available in restriction, TOPO, and Gateway cloning versions and as untagged and epitope-tagged versions, whereas the pcDNA3.3-TOPO vector is a TOPO TA-adapted, untagged vector that can be used to express native proteins without extraneous amino acids, and is hence ideal for antibody production and structural biology.

Do you offer a GFP-expressing mammalian expression vector that I can use as a control to monitor my transfection and expression?

We offer pJTI R4 Exp CMV EmGFP pA Vector, Cat. No. A14146, which you can use to monitor your transfection and expression.

I am working with a mouse cell line and would like to express my gene at high levels using one of your vectors with the CMV promoter. Do you foresee any problems with this approach?

The CMV promoter is known to be downregulated over time in mouse cell lines. Hence, we recommend using one of our non-CMV vectors, such as those with the EF1alpha or UbC promoter, for long-term expression in mouse cell lines.

Do I need to include a consensus Kozak sequence when I clone my gene of interest into one of your mammalian expression vectors?

The consensus Kozak sequence is A/G NNATGG, where the ATG indicates the initiation codon. Point mutations in the nucleotides surrounding the ATG have been shown to modulate translation efficiency. Although we make a general recommendation to include a Kozak consensus sequence, the necessity depends on the gene of interest and often, the ATG alone may be sufficient for efficient translation initiation. The best advice is to keep the native start site found in the cDNA unless one knows that it is not functionally ideal. If concerned about expression, it is advisable to test two constructs, one with the native start site and the other with a consensus Kozak. In general, all expression vectors that have an N-terminal fusion will already have an initiation site for translation.

Find additional tips, troubleshooting help, and resources within our Protein Expression Support Center.

Do I need to include a ribosomal binding site (RBS/Shine Dalgarno sequence) or Kozak sequence when I clone my gene of interest?

ATG is often sufficient for efficient translation initiation although it depends upon the gene of interest. The best advice is to keep the native start site found in the cDNA unless one knows that it is not functionally ideal. If concerned about expression, it is advisable to test two constructs, one with the native start site and the other with a Shine Dalgarno sequence/RBS or consensus Kozak sequence (ACCAUGG), as the case may be. In general, all expression vectors that have an N-terminal fusion will already have a RBS or initiation site for translation.

Find additional tips, troubleshooting help, and resources within our Protein Expression Support Center.

What is the best molar ratio of PCR product:vector to use for TOPO TA cloning? Is there an equation to calculate the quantity to use?

We suggest starting with a molar ratio of 1:1 (insert:vector), with a range of 0.5:1 to 2:1. The quantity used in a TOPO cloning reaction is typically 5-10 ng of a 2 kb PCR product.

Equation:

length of insert (bp)/length of vector (bp) x ng of vector = ng of insert needed for 1:1 (insert:vector ratio)

What is the best ratio of insert:vector to use for cloning? Is there an equation to calculate this?

The optimal ratio is 1:1 insert to vector. Optimization can be done using a ratio of 0.5-2 molecules of insert for every molecule of the vector.

Equation:

length of insert (bp)/length of vector (bp) x ng of vector = ng of insert needed for 1:1 insert:vector ratio

Does Platinum Taq DNA Polymerase High Fidelity enzyme mix leave 3' A-overhangs on the PCR product for subsequent cloning into a TOPO TA or original TA vector?

Yes, the enzyme mix leaves 3' A-overhangs on a portion of the PCR products. However, the cloning efficiency is greatly decreased compared to that obtained with Taq polymerase alone. It is recommended to add 3' A-overhangs to the product for TA cloning.

I'm seeing a lot of vector-only colonies when I try to perform a negative control reaction using vector only (no insert) for a TOPO reaction. Is my TOPO vector re-ligating?

Using the vector only for transformation is not a recommended negative control. The process of TOPO-adaptation is not a 100% process, therefore, there will be “vector only” present in your mix, and colonies will be obtained.

I'm trying to clone in my phosphorylated PCR product into a TOPO vector, and I'm getting no colonies. However, when I clone the same product into a TA vector, everything works perfectly. Why is this?

Phosphorylated products can be TA cloned but not TOPO cloned. This is because the necessary phosphate group is contained within the topoisomerase-DNA intermediate complex of the vector. TOPO vectors have a 3' phosphate to which topoisomerase is covalently bound and a 5' phosphate. Non-TOPO linear vectors (TA and Blunt) have a 3' OH and a 5' phosphate. Phosphorylated products should be phosphatased (CIP) before TOPO cloning.

I'm able to get a lot of colonies, however, none contain my insert of interest. What should I do?

You may be cloning in an artifact. TA and TOPO Cloning are very efficient for small fragments (< 100 bp) present in certain PCR reactions. Gel-purify your PCR product using either a silica-based DNA purification system or electroelution. Be sure that all solutions are free of nucleases (avoid communal ethidium bromide baths, for example.)

A majority of colonies are blue or light blue, with very few white colonies. What should I do?

There could be a few possibilities for this:

- The insert does not interrupt the reading frame of the lacZ gene. If you have a small insert (< 500 bp), you may have light blue colonies. Analyze some of these blue colonies as they may contain insert.
- A polymerase that does not add 3' A-overhangs was used. If you used a proofreading enzyme, you will need to do a post-reaction treatment with Taq polymerase to add the 3' A-overhangs.
- PCR products were gel-purified before ligation. Gel purification can remove the single 3' A- overhangs. Otherwise, optimization of your PCR can be performed so that you can go directly from PCR to cloning.
- The PCR products were stored for a long period of time before ligation reaction. Use fresh PCR products. Efficiencies are reduced after as little as 1 day of storage.
- Too much of the amplification reaction was added to the ligation. The high salt content of PCR can inhibit ligation. Use no more than 2-3 µl of the PCR mixture in the ligation reaction.
- The molar ratio of vector:insert in the ligation reaction may be incorrect. Estimate the concentration of the PCR product. Set up the ligation reaction with a 1:1 or 1:3 vector:insert molar ratio.
On a typical plate there are a few white colonies which do not contain insert. These are usually larger than the other colonies and are due to a deletion of a portion of the plasmid sequence by a rare recombination event (usually from the polylinker to a site in the F1 origin). To find a colony with an insert it is best to pick clones of a variety of color and pattern for analysis. Often an insert will generate two distinct patterns according to its orientation.

I'm getting no colonies after transformation. What should I do?

No colonies may occur due to the following problems:

Bacteria were not competent. Use the pUC18 vector included with the One Shot module to check the transformation efficiency of the cells.
- Incorrect concentration of antibiotic on plates, or the plates are too old. Use 100 µg/mL of ampicillin or 50 µg/mL kanamycin. Be sure ampicillin plates are fresh (< 1 month old).
- The product was phosphorylated (TOPO cloning only). Phosphorylated products can be TA-cloned but not TOPO-cloned. This is because the necessary phosphate group is contained within the topoisomerase-DNA intermediate complex of the vector. The TOPO vector has a 3' phosphate to which topoisomerase is covalently bound and a 5' phosphate. The non- TOPO vectors (TA and Blunt) have a 3' OH and a 5' phosphate. Phosphorylated products should be phosphatased (CIP) before TOPO-cloning.

I'm getting low cloning efficiency with my TOPO cloning reactions. What should I do?

Please consider the following possible causes:
- pH > 9: Check the pH of the PCR amplification reaction and adjust with 1 M Tris-HCl, pH 8.
- Excess (or overly dilute) PCR product: Reduce (or concentrate) the amount of PCR product.
- Incomplete extension during PCR: Be sure to include a final extension step of 7 to 30 minutes during PCR. Longer PCR products will need a longer extension time.
- Cloning large inserts (>1 kb): Try one or all of the following suggestions: Increase amount of insert. Incubate the TOPO cloning reaction longer. Gel-purify the insert using either a silica-based DNA purification system (e.g., PureLink system) or electroelution. Be sure that all solutions are free of nucleases (avoid communal ethidium bromide baths, for example.)
- PCR product does not contain sufficient 3' A-overhangs even though you used Taq polymerase: Increase the final extension time to ensure all 3' ends are adenylated. Taq polymerase is less efficient at adding a nontemplate 3' A next to another A. Taq is most efficient at adding a nontemplate 3' A next to a C. You may have to redesign your primers so that they contain a 5' G instead of a 5´ T.

I'm getting very few colonies after transformation of my TOPO cloning reaction. How can I increase the number of primary colonies?

Please try the suggestions below to increase the number of colonies.
- Longer incubation of the TOPO cloning reaction at room temperature, provided that the 6X Salt solution is added to the reaction.
- Electroporation can give significant increases in colony numbers; often 10-20 fold higher. However, if doing electroporation, it is important that the TOPO reaction mix contains diluted Salt solution or, for best results, precipitated prior to transformation. For high primary transformants by electroporation it is recommended to:
- Add 100 µL double diH2O to the 6 µL TOPO reaction and incubate 10 more minutes at 37 degrees C.
- Precipitate by adding 10 µL 3 M Na-Acetate, 2 µL 20 µg/µL glycogen, 300 µL 100% ethanol. Place on dry ice or –80 degrees C for 20 min, spin at top speed in a microcentrifuge at 4 degrees C for 15 min. Wash pellet with 800 µL 80% ethanol, spin at top speed for 10 min, pour off ethanol, spin 1 min, and remove remaining ethanol without disturbing pellet. Dry pellet (air-dry or speed-vac).
- Resuspend pellet in 10 µL ddH2O and electroporate 3.3 µL of resuspended DNA according to a normal electroporation protocol. This electroporation protocol can yield up to 20 fold more colonies than chemical transformation of an equivalent TOPO-reaction. The addition of the 100 µL ddH2O followed by 10 mins incubation is not absolutely necessary, but it sufficiently dilutes the reaction and may help inactivate topoisomerase so that it is more easily electroporated.

I'm planning on cloning a 1kb fragment for sequencing and want to minimize the amount of vector sequence in my data. Which of your vectors should I use?

We would suggest using our TOPO TA cloning kit for sequencing, which contains the pCR 4 TOPO vector, or our Zero Blunt TOPO PCR cloning kit for sequencing, which contains the pCR4Blunt-TOPO vector.

I'm trying to decide between your pCR2.1 TOPO and pCR4-TOPO vectors to clone a 150 bp PCR product for sequencing. Which would you recommend?

Due to the small size of your product, we recommend using the pCR 2.1 TOPO vector for your cloning. This size fragment would not be able to fully interrupt the ccdB gene in the pCR4-TOPO vector, and therefore, you may not get colonies as ccdB is lethal to E. coli.

What are the insert size limitations of TOPO cloning kits?

Regular TOPO TA Cloning kits are efficient for cloning PCR products up to approximately 2-3 kb. With PCR products larger than 3 kb, the efficiency of cloning drops significantly. The TOPO XL PCR Cloning Kit has been optimized for TOPO cloning of long (3-10 kb) PCR products.

If using the regular TOPO kits, here are some tips to improve efficiency:

1. Use crystal violet instead of ethidium bromide (EtBr) to visualize the PCR for gel isolation to avoid DNA nicks
2. Increase incubation time of the TOPO reaction to 30 mins
3. Keep insert:vector molar ratio low, optimally 1:1
4. Dilute reaction to 20 µL, while maintaining same amount of vector and insert. Increase the volume of the salt solution to 3.7 µL to compensate for the increase in volume. Diluting the reaction reduces the competition for the vector ends.

Can I store my TOPO vector plus insert reaction? At what temperature?

Storage of the TOPO vector plus insert reaction for 1 week at 4 degrees C has shown no detectable decrease in the cloning efficiency of the TOPO reaction, as >95% of the colonies have insert. However, the total number of colonies was decreased by 10-fold. Storage of the TOPO reaction mix overnight at 4 degrees C showed little to no decrease in the number of colonies when compared to fresh TOPO reaction mix.

What is the difference between a stop solution and salt solution? What is its function in the TOPO kit?

The composition of the 6X Stop solution is 0.3 M NaCl, 0.06 M MgCl2, and the composition of the 6X Salt solution is 1.2 M NaCl, 0.06 M MgCl2. Stop solution is only included in the TOPO XL Cloning kit whereas Salt solution is currently included in all of the other TOPO cloning kits. These solutions prevent free topoisomerase from re-binding and nicking the plasmid, which would reduce the number of colonies from a TOPO reaction.

What can inhibit the TOPO cloning reaction?

When doing a TOPO cloning reaction, 2 µl of a PCR reaction containing up to 10% DMSO or 1.3 M betaine will not interfere with the TOPO reaction. Formamide and high levels of glycerol will inhibit the reaction. These reagents are usually added to the PCR reaction to enhance the yield of the PCR product, e.g., to reduce the effect of secondary structure or assist in amplification of GC-rich sequences. The effects of tricine or acetamide have not been tested on the TOPO cloning reaction.

What considerations should I take into account when designing primers for PCR of an insert which will be cloned into a TOPO vector?

PCR primers should not have 5'-phosphates when cloning into any TOPO vector, as the presence of 5'-phosphates inhibit the TOPO cloning reaction. Phosphorylated products can be TA-cloned but not TOPO-cloned. This is because the necessary phosphate group is contained within the topoisomerase-DNA intermediate complex of the vector. TOPO vectors have a 3' phosphate to which topoisomerase is covalently bound and a 5' phosphate. Non-TOPO linear vectors (TA and Blunt) have a 3' OH and a 5' phosphate. Phosphorylated products should be treated with phosphatase (CIAP) prior to TOPO-cloning. Treatment with CIAP may raise efficiency to 25%. PCR products generated with 5'-biotinylated primers (or any other 5'-label including 5'-Cy5) will not ligate into any of the TOPO vectors due to steric hindrance.

Do I need to gel purify my PCR product for TOPO cloning?

Gel purification is not required if the gel indicates that the PCR product is clean with no visible non-specific bands or primer dimers. It is recommended if the PCR product is >1.5 kb or if non-specific bands and primer dimers are visible on the gel. Smaller products clone much more efficiently into the vector than larger products; therefore, they should be eliminated from the sample prior to cloning. There is some reduction in A-overhangs if the PCR product is gel purified, which along with PCR product loss during the procedure may slightly reduce total number of colonies. However, the percentage of colonies with insert does not change; it is typically >90% recombinant clones.

I typically store my PCR products before TOPO cloning. Is this okay?

For optimal TOPO cloning, we recommend using fresh PCR products.

What are the advantages of using a TOPO TA cloning system compared to traditional TA cloning?

TA cloning ligates the insert and vector using a T4 DNA ligase, while TOPO TA cloning uses the intrinsic properties of topoisomerase, which ligates the insert and vector during a 5 minute desktop reaction. TOPO TA cloning results in >95% recombinants, while TA cloning results in >80% recombinants.

How do I adapt my cloning vector for TOPO cloning?

We offer a custom service for TOPO cloning adaptation services. Our scientists can prepare your vector for either blunt TOPO cloning, TOPO TA cloning, or directional TOPO cloning of PCR products.

Can I order my TOPO vector as a standalone product? I have plenty of competent cells.

Yes, our pCR.1 TOPO TA (Cat. No. 450641), pCR4-TOPO TA (Cat. No. 450030), pCRBluntII-TOPO (Cat. No. 450245) are available separately.

Can I run the TOPO vector on a gel?

No, we do not recommend this as these vectors contain the topoisomerase DNA protein complex conjugated to the end of the vector.

What range of PCR product (molar ratios and ng quantities) do you suggest for TOPO TA cloning?

We suggest starting with a molar ratio of 1:1 (insert:vector), with a range of 0.5:1 to 2:1 (insert:vector). The ng quantities should be between 5-10 ng of a 2 kb PCR product in a TOPO cloning reaction.

What are some of the prerequisites for TOPO cloning?

Please consider the following before TOPO cloning:

- TOPO cloning cannot ligate DNA with a 5' phosphate group.
- TOPO cloning will decrease in efficiency inversely with the size of the insert (above 3 kb) unless using the TOPO XL cloning kit.
- TOPO vectors contain different antibiotic resistance markers which should be considered before purchase.
- TOPO TA vectors accept fragments containing a 3' A overhang while Zero Blunt vectors accept fragments that are blunt-ended.

I received my TOPO vector and the solution is colored. Is it okay to use?

TOPO and TOPO TA vectors (non-directional) have phenol red dye added. The color should be pink (or yellow) at room temperature. If it turns blue when PCR product is added, the PCR product buffer is too basic and the number of transformed colonies will drop. When the solution is yellow, it signifies an acidic pH. At a pH 2.0, TOPO vectors still maintain high cloning efficiency. Directional TOPO and Zero Blunt TOPO vectors have bromophenol blue dye added.

I have a TOPO TA Cloning kit with TOP10 cells. I ran out of competent cells but still have vector left. I also have subcloning DH5? cells and TOP10F' cells in the freezer. Are either of these cells compatible? What strain features should I be aware of?

Subcloning DH5? cells are a compatible strain, but you will get lower efficiency (10e6 vs 10e9) and therefore risk getting fewer clones. Top10F' is also compatible, but if blue/white screening is performed, IPTG along with X-gal will be needed for detection due to the expression of the lacIq repressor present in cells containing an F' episome.

I'm getting overgrowth of colonies. Why?

Ensure that you are using the correct antibiotic at the appropriate concentration. Additionally, make sure the antibiotic is not expired. If colonies exhibit unexpected morphologies, contamination could be a factor. Check your S.O.C. medium and LB growth medium.

I'm only getting white colonies, but none of the clones have an insert. What can I do?

Here are a few suggestions:

- Small fragments/linkers are cloning in to your vector instead of your insert; to correct this, gel-purify the insert before ligation
- Ensure that the correct concentrations of X-gal and/or IPTG (if vector contains the lacIq marker) are used
- If spreading X-gal and/or IPTG on your plate, allow sufficient time for the reagents to diffuse into the plate
- Incubate your plate for a longer period to ensure full color development

I'm trying to transform large plasmid, 40 kb in size. What strain should I use?

While there is no specific strain that works better with large plasmids, it is important to focus on transformation efficiency. For larger plasmids, chemically competent cells with highest efficiency are suggested, such as OmniMAX 2, TOP10, etc. We would recommend using an electrocompetent cell strain with plasmids larger than 20 kb for best efficiency.

I'm trying to clone a gene that has multiple repeated sequences into my pCR2.1-TOPO vector, followed by transformation into TOP10 cells. My clones contain random rearrangements and deletions. What can I do?

With any strain, the first thing to try would be to lower the growth temperature of the culture to 30 degrees C or even lower (room temperature). Slower growth will generally allow E. coli to tolerate difficult sequences better. If reducing the growth temperature doesn't help, you may want to consider using a competent cell strain such as Stbl2 or Stbl4 cells, which have been shown to accommodate this type of sequence better than other strains in the same conditions.

I'm getting no colonies at all on my plates. Can you help?

We recommend trying the following:
- Carry out the puc19 transformation control; this gives you information about the performance of the cells.
- Check plates for expiration and correct media used (LB/agar).
- Confirm that the correct antibiotic and concentration was used.

I'm transforming pCR2.1-TOPO clones into TOP10F' cells. Will I need to add IPTG to my plates along with X-gal for blue/white screening? What if I used TOP10 cells instead?

The F' episome in TOP10F' has a lacIq marker, which over-expresses the lac repressor. IPTG must be added to LB plates along with X-gal to see beta-galactosidase expression and blue color in this strain. TOP10, on the other hand, does not require IPTG for blue/white screening.

I'm plating my untransformed TOP10 cells on ampicillin as a negative control, but see a lot of colonies on the plates.

There are a few conditions that can lead to this: SOC medium or other media used when plating was contaminated, DNA was contaminated with amp-resistant microbes, you used old plates with degraded amp, or the competent cells themselves were contaminated.

I'm subcloning fragments of yeast genomic DNA into a TOPO vector. I'm seeing a lot of deletions in the clones I'm selecting. What can I do?

If you are using a mcr/mrr(+) competent cell strain, cellular enzymes may be recognizing eukaryotic methylation patterns on the yeast genomic DNA and deleting or rearranging it. Try a mcr/mr(-) strain such as Top10, DH10B, or OmniMAX 2.

I've cloned my gene into the pCR2.1-TOPO vector and transformed into the TOP10 cells that came with the kit. I then did a plasmid miniprep followed by digestion of the DNA with XbaI. However, the vector is not cutting correctly. What happened?

XbaI cutting site is a Dam-methylation sensitive restriction site. TOP10 is a dam(+) strain, which means it expresses the methylating enzyme, Dam. You can try re-transforming into a dam(-) strain, such as INV110. Other dam- (and dcm-) sensitive restriction sites include the following:

- Dam: Bcl I, Cla I, Hph I, Mbo I, Mbo II, Taq I, Xba I, BspH I, Nde II, Nru I
- Dcm: Ava II, EcoO 109 I, EcoR II, Sau96 I, ScrF, Stu I, Aat I, Apa I, Bal I, Kpn I, ISfi I

What suggestions can you make for blue/white screening?

1. Use pUC or pUC-based vectors that contain the portion of the lacZ gene that allows for ? complementation.
2. Select an E. coli strain that carries the lacZdeltaM15 marker.
3. Plate transformations on plates containing X-gal. Spread 50 µg of 2% X-gal or 100 microliters of 2% bluo-gal (both can be dissolved in DMF or 50:50 mixture of DMSO:water) on the surface of a 100 mm plate and let dry. Alternatively, add directly to the cooled medium (~50 degrees C) before pouring the plates at a final concentration of 50 µg/mL for X-gal and 300 µg/mL for bluo-gal. Plates are stable for 4 weeks at 4 degrees C.
4. If the strain used carries the lacIq marker, add IPTG to induce the lac promoter. Spread 30 µl of 100 mM IPTG (in water) on 100 mm plates. Alternatively, add the IPTG directly to cooled medium (~50 degrees C) before pouring the plates to a final concentration of 1 mM. Plates are stable for 4 weeks at 4 degrees C.
5. Do not plate E. coli on medium containing glucose if using X-gal or bluo-gal for blue-white screening. Glucose competes as a substrate and prevents cells from turning blue.

I want to store my transformed cells long term. Do you have a protocol for this?

For long-term storage, preparation of glycerol stocks stored at -70 degrees C is recommended. Follow the protocol below:
1. Pick one colony into 5 mL LB broth or S.O.C. medium. Grow overnight at 37 degrees C.
2. Prepare glycerol solution: 6 mL of S.O.B. medium and 4 mL of glycerol.
3. Take one volume of cells and add one volume of glycerol solution and mix.
4. Freeze in ethanol/dry ice. Store at -70 degrees C.

Can I transform 2 plasmids into the same cell?

Yes, this is possible. We recommend using saturating amounts of DNA (10 ng of each plasmid). Make sure that the origin of replication is different in each plasmid so that they can both be maintained in the cell. If the ori is the same, the plasmids will compete for replication and the one with even a slight disadvantage will be lost. Alternatively, cells with a resident plasmid can be electroporated with a second plasmid without “electrocuring” taking place.

What concentrations do you typically recommend for X-gal and IPTG for blue/white screening?

In plates, we recommend 50 µg/mL X-gal and 1 mM IPTG (0.24 mg/mL). When spreading directly onto agar plates, we recommend 40-50 µl of 40 mg/mL X-gal (2% stock) in dimethylformamide and 30-40 µl of 100 mM IPTG on top of the agar. Let the X-gal and IPTG diffuse into the agar for approximately 1 hour. Do not plate on media containing glucose, as it competes with X-gal or bluo-gal and prevents cells from turning blue.

How is competent cell efficiency measured? How is it calculated?

Competent cell efficiency is measured by transformation efficiency. Transformation efficiency is equal to the number of transformants, or colony forming units, per microgram of plasmid DNA (cfu/microgram).

What are some tips you can give me to obtain the highest transformation efficiency with my competent cells?

Some suggestions that will help you to obtain the highest transformation efficiency are:
- Thaw competent cells on ice instead of room temperature; do not vortex cells.
- Add DNA to competent cells once thawed.
- Ensure that the incubation times are followed as outlined in the competent cell protocol for the strain you are working with; changes in the length of time can decrease efficiency.
- Remove salts and other contaminants from your DNA sample; DNA can be purified before transformation using a spin column, or phenol/chloroform extraction and ethanol precipitation can be employed.

I'm trying to decide between the TOP10, DH5?, and Mach1 strains you have for my TOPO TA Cloning reactions. Can you explain the significant differences between these strains?

DH5? cells are commonly used for routine cloning, but are mcr/mrr+, and therefore not recommended for genomic cloning. The TOP10 competent cells, on the other hand, contain mutated mcr/mrr, and therefore are a good choice for routine cloning and can be used for cloning of methylated DNA, such as eukaryotic genomic DNA. Our Mach1 strain is the fastest growing cloning strain that is T1 phage resistant.

I see small satellite colonies on my LB+Amp plates. Why is this?

These small colonies are most likely caused by degradation of the Ampicillin. The colonies are just untransformed cells that grow on LB with degraded Amp. In order to circumvent this scenario, you can try to:
1. Plate cells at a lower density
2. Use fresh LB-Amp plates or replace Ampicillin with carbenicillin.
3. The plates should not be incubated for more than 20 hours at 37 degrees C. Beta-lactamase, the enzyme produced from the Ampicillin-resistance gene, is secreted from the Amp-resistant transformants and inactivates the antibiotic in the area surrounding the transformant colony. This inactivation of the selection agent allows satellite colonies (which are not truly Amp-resistant) to grow. This is also true if carbenicillin is being used.

I'm able to see colonies on a plate, but when I pick them for liquid culture, no growth is observed. Why?

One possible explanation could be toxicity associated with the insert. This toxicity does not affect slow growing cells on solid medium but is much stronger in faster growth conditions like liquid medium.

Suggestions:
1. Use TOP10F' or any other strain with the LacIq repressor
2. Try using any other strain appropriate for cloning.
3. Lower growth temperature to 27 - 30 degrees C and grow the culture longer
4. Another possibility to explain lack of growth is possible phage contamination. In this situation we recommend using an E. coli strain that is T1 phage resistant like DH5alpha-T1R.

The clones I'm selecting show deleted inserts. Why?

This may be caused by the instability of the insert DNA in TOP10 E. Coli. In this case, E.coli strains such as Stbl2, Stbl3, or Stbl4 have been shown to support the propagation of DNA with multiple repeats, retroviral sequences, and DNA with high GC content better than other strains.

I'm getting low to no colonies after transformation. Why?

Some possible causes and remedies are:
- Ligase function is poor. Check the age of the ligase and function of the buffer.
- Competent cells are not transforming. Test the efficiency of the cells with a control supercoiled vector, such as puc19.
- Both molecules were de-phosphorylated.
- Inhibition of ligation by restriction enzymes and residual buffer. Try transformation of uncut vector, clean up restriction with phenol, or carry out PCR cleanup/gel extraction before ligation.
- Incorrect antibiotic selection used. Check the plasmid and plates and make sure concentration of antibiotic used is correct.

If nothing above applies, low to no colonies may be due to instability of the insert DNA in your competent cells. In this case, E. coli strains such as Stbl2, Stbl3, or Stbl4 have been shown to support the propagation of DNA with multiple repeats, retroviral sequences, and DNA with high GC content better than other strains.

How does selection with the LacZ gene work?

If working with a vector that contains the lac promoter and the LacZ ? fragment (for ? complementation), blue/white screening can be used as a tool to select for presence of the insert. X-gal is added to the plate as a substrate for the LacZ enzyme and must always be present for blue/white screening. The minimum insert size needed to completely disrupt the lacZ gene is >400 bp. If the LacIq repressor is present (either provided by the host cells, for example TOP10F', or expressed from the plasmid), it will repress expression from the lac promoter thus preventing blue/white screening. Hence, in the presence of the LacIq repressor, IPTG must be provided to inhibit the LacIq. Inhibition of LacIq permits expression from the lac promoter for blue/white screening.

How does ccdB selection work?

TOPO vectors containing the LacZ-ccdB cassette allow direct selection of recombinants via disruption of the lethal E. coli gene, ccdB. Ligation of a PCR product disrupts expression of the LacZ-ccdB gene fusion permitting growth of only positive recombinants upon transformation. Cells that contain non-recombinant vector are killed upon plating. Therefore, blue/white screening is not required. When doing blue/white color screening of clones in TOPO vectors containing the LacZ-ccdB cassette, colonies showing different shades of blue may be observed. It is our experience that those colonies that are light blue as well as those that are white generally contain inserts. The light blue is most likely due to some transcription initiation in the presence of the insert for the production of the lacZ alpha without enough ccdB expressed to kill the cells and is insert dependent. To completely interrupt the lacZ gene, inserts must be >400 bp; therefore an insert of 300 bp can produce a light blue colony. A white colony that does not contain an insert is generally due to a spontaneous mutation in the ccdB gene.
A minimum insertion of 150 bp is needed in order to ensure disruption of the ccdB gene and prevent cell death. (Reference: Bernard et al., 1994. Positive-selection vectors using the F plasmid ccdB killer gene. Gene 148: 71-74.) Strains that contain an F plasmid, such as TOP10F', are not recommended for transformation and selection of recombinant clones in any TOPO vector containing the ccdB gene. The F plasmid encodes the CcdA protein, which acts as an inhibitor of the CcdB gyrase-toxin protein. The ccdB gene is also found in the ccd (control of cell death) locus on the F plasmid. This locus contains two genes, ccdA and ccdB, which encode proteins of 72 and 101 amino acids respectively. The ccd locus participates in stable maintenance of F plasmid by post-segregational killing of cells that do not contain the F plasmid. The CcdB protein is a potent cell-killing protein when the CcdA protein does not inhibit its action.

How does blue/white screening work?

If working with a vector that contains the lac promoter and the LacZ alpha fragment (for ? complementation), blue/white screening can be used as a tool to select for presence of the insert. X-gal is added to the plate as a substrate for the LacZ enzyme and must always be present for blue/white screening. The minimum insert size needed to completely disrupt the lacZ gene is >400 bp. If the LacIq repressor is present (either provided by the host cells, for example TOP10F', or expressed from the plasmid) it will repress expression from the lac promoter, thus preventing blue/white screening. Hence in the presence of the LacIq repressor, IPTG must be provided to inhibit the LacIq. Inhibition of LacIq permits expression from the lac promoter for blue/white screening. X-gal (also known as 5-bromo-4-chloro-3-indolyl β-D-glucopyranoside) is soluble in DMSO or DMF, and can be stored in solution in the freezer for up to 6 months. Protect the solution from light. Final concentration of X-gal and IPTG in agar plates: Prior to pouring plates, add X-gal to 20 mg/mL and IPTG to 0.1 mM to the medium. When adding directly on the surface of the plate, add 40 µl X-gal (20 mg/mL stock) and 4 µl IPTG (200 mg/mL stock).

Can I use TOPOTA pCR2.1 or pCR II or pCR4 for my protein expression experiments?

No, these vectors do not contain a functional promoter to express your gene of interest. These vectors are typically for subcloning or sequencing.

Which PCR polymerases do you recommend for TA/Blunt/D-TOPO cloning and why?

TA Cloning:
- This cloning method was designed for use with pure Taq polymerases (native, recombinant, hot start); however, High Fidelity or Taq blends generally work well with TA cloning. A 10:1 or 15:1 ratio of Taq to proofreader polymerase will still generate enough 3' A overhangs for TA cloning.
- Recommended polymerases include Platinum Taq, Accuprime Taq, Platinum or Accuprime Taq High Fidelity, AmpliTaq, AmpliTaq Gold, or AmpliTaq Gold 360.

Blunt cloning:
- Use a proofreading enzyme such as Platinum SuperFi DNA Polymerase.

Directional TOPO cloning:
- Platinum SuperFi DNA Polymerase works well.

What PCR enzyme would you recommend for use with the Directional TOPO Cloning Kits?

For the Directional TOPO Cloning Vectors, a PCR product must be generated by a proofreading enzyme to create a blunt product. Pfx50 or Accuprime Pfx and Accuprime Pfx Supermix from Thermo Fisher Scientific are recommended for use.

When cloning a Pfx-amplified PCR product, the insert to vector ratio is an important consideration. The PCR product generally needs to be diluted since Pfx generates a high concentration of product and using too much insert DNA can hamper the TOPO reaction. A 1:1 molar ratio of vector to insert (or about 2-10ng of insert) is recommended.

Can I use a DNA polymerase mixture containing both Taq polymerase and a proofreading polymerase for TA Cloning?

If you wish to use a polymerase mixture containing Taq polymerase and a proofreading polymerase, Taq must be used in excess with a 10:1 ratio of Taq to the proofreading enzyme to ensure the presence of 3´ A-overhangs on the PCR product. If you use polymerase mixtures that do not have enough Taq polymerase or a proofreading polymerase only, you can add 3' A-overhangs following PCR. See the vector product manuals for details.

Some examples of Taq blends that are compatible with TOPO TA Cloning are Platinum Taq DNA Polymerase High Fidelity and AccuPrime Taq DNA Polymerase High Fidelity.

How does TA Cloning work?

Taq polymerase has a non-template-dependent terminal transferase activity that adds a single deoxyadenosine (A) to the 3´ ends of PCR products. The linearized vector supplied in our TA Cloning kits have single, overhanging 3´ deoxythymidine (T) residues. This allows PCR inserts to ligate efficiently with the vector.

What are the melting temperatures for the M13 Forward (-20) and M13 Reverse primers in the TOPO Cloning and Zero Blunt Kits?

Assuming that the primer is at a 50 nM final concentration and 50 mM final salt concentration, the melting temperatures are: M13 Forward (-20) Primer = 52.7 and the M13 Reverse Primer = 45.3. For use in the control PCR reaction we recommend using an annealing temperature of 56C.

Can you tell me the difference between a Shine-Dalgarno sequence and a Kozak sequence?

Prokaryotic mRNAs contain a Shine-Dalgarno sequence, also known as a ribosome binding site (RBS), which is composed of the polypurine sequence AGGAGG located just 5’ of the AUG initiation codon. This sequence allows the message to bind efficiently to the ribosome due to its complementarity with the 3’-end of the 16S rRNA. Similarly, eukaryotic (and specifically mammalian) mRNA also contains sequence information important for efficient translation. However, this sequence, termed a Kozak sequence, is not a true ribosome binding site, but rather a translation initiation enhancer. The Kozak consensus sequence is ACCAUGG, where AUG is the initiation codon. A purine (A/G) in position -3 has a dominant effect; with a pyrimidine (C/T) in position -3, translation becomes more sensitive to changes in positions -1, -2, and +4. Expression levels can be reduced up to 95% when the -3 position is changed from a purine to pyrimidine. The +4 position has less influence on expression levels where approximately 50% reduction is seen. See the following references:

- Kozak, M. (1986) Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 44, 283-292.
- Kozak, M. (1987) At least six nucleotides preceding the AUG initiator codon enhance translation in mammalian cells. J. Mol. Biol. 196, 947-950.
- Kozak, M. (1987) An analysis of 5´-noncoding sequences from 699 vertebrate messenger RNAs. Nucleic Acids Res. 15, 8125-8148.
- Kozak, M. (1989) The scanning model for translation: An update. J. Cell Biol. 108, 229-241.
- Kozak, M. (1990) Evaluation of the fidelity of initiation of translation in reticulocyte lysates from commercial sources. Nucleic Acids Res. 18, 2828.

Note: The optimal Kozak sequence for Drosophila differs slightly, and yeast do not follow this rule at all. See the following references:

- Romanos, M.A., Scorer, C.A., Clare, J.J. (1992) Foreign gene expression in yeast: a review. Yeast 8, 423-488.
- Cavaneer, D.R. (1987) Comparison of the consensus sequence flanking translational start sites in Drosophila and vertebrates. Nucleic Acids Res. 15, 1353-1361.

Find additional tips, troubleshooting help, and resources within our Protein Expression Support Center.

I sequenced one of your vectors after PCR amplification and observed a difference from what is provided online (or in the manual). Should I be concerned?

Our vectors have not been completely sequenced. Your sequence data may differ when compared to what is provided. Known mutations that do not affect the function of the vector are annotated in public databases.

Are your vectors routinely sequenced?

No, our vectors are not routinely sequenced. Quality control and release criteria utilize other methods.

How was the reference sequence for your vectors created?

Sequences provided for our vectors have been compiled from information in sequence databases, published sequences, and other sources.

What is the consensus Kozak sequence and what is the function of the Kozak sequence?

Eukaryotic (and specifically mammalian) mRNA contains sequence information that is important for efficient translation. However, this sequence, termed a Kozak sequence, is not a true ribosome binding site, but rather a translation initiation enhancer. The Kozak consensus sequence is ACCAUGG, where AUG is the initiation codon. A purine (A/G) in position -3 has a dominant effect; with a pyrimidine (C/T) in position -3, translation becomes more sensitive to changes in positions -1, -2, and +4. Expression levels can be reduced up to 95% when the -3 position is changed from a purine to pyrimidine. The +4 position has less influence on expression levels where approximately 50% reduction is seen. See the following references:

Kozak, M. (1986) Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 44, 283-292.
Kozak, M. (1987) At least six nucleotides preceding the AUG initiator codon enhance translation in mammalian cells. J. Mol. Biol. 196, 947-950.
Kozak, M. (1987) An analysis of 5´-noncoding sequences from 699 vertebrate messenger RNAs. Nucleic Acids Res. 15, 8125-8148.
Kozak, M. (1989) The scanning model for translation: An update. J. Cell Biol. 108, 229-241.
Kozak, M. (1990) Evaluation of the fidelity of initiation of translation in reticulocyte lysates from commercial sources. Nucleic Acids Res. 18, 2828.

Note: The optimal Kozak sequence for Drosophila differs slightly, and yeast do not follow this rule at all. See the following references:

Romanos, M.A., Scorer, C.A., Clare, J.J. (1992) Foreign gene expression in yeast: a review. Yeast 8, 423-488.
Cavaneer, D.R. (1987) Comparison of the consensus sequence flanking translational start sites in Drosophila and vertebrates. Nucleic Acids Res. 15, 1353-1361.

Find additional tips, troubleshooting help, and resources within our Protein Expression Support Center.

Do you offer pcDNA 3.3-TOPO TA Cloning Kit (Cat. No. K830001)?

pcDNA 3.3-TOPO TA Cloning Kit (Cat. No. K830001) has been discontinued. As an alternative, you may use pcDNA 3.4 TOPO TA Cloning Kit (Cat. No. A14697).

Find additional tips, troubleshooting help, and resources within our Protein Expression Support Center.