Topics: AccuPOL, AMV One Tube Hot Start RT-PCR Kit, BL21 (DE3) Competent Cells, BL21 (DE3) pLysS Competent Cells, Buffer Optimization Kit, cDNA Direct, dNTP Sets, GC10 High Efficiency Competent Cells (comparable to DH10B), SuperPath GC10 ElectroCompetent Cells, GC5 High Efficiency Competent Cells (comparable to DH5alpha), GC5 Value Efficiency Competent Cells (comparable to DH5alpha), Rapid Ligation Kit, Taq with Ammonium Buffer, Taq with Magnesium Free Buffer, Taq DNA Pol 1.1X Master Mix Kit, TEMPase, UniPOL.
AccuPOL
Q: Does the enzyme leave A overhangs?
A: No. AccuPOL is a proofreader which produces blunt ends.
Q: How much enzyme should be used per reaction?
A: 2.5 units is standard. We suggest trying different amounts to optimize your conditions.
Q: What is the optimal extension temperature?
A: The optimal extension temperature is 72 degrees C, same as for Taq.
Q: Would you suggest AccuPOL for cloning?
A: You can generate your fragment with AccuPOL and use it for blunt-end cloning. If you want to clone the AccuPOL amplified fragment in a T/A cloning vector, you have to incubate your fragment with Taq Polymerase for 10 min at 72 C.
Q: What concentration of dNTPs should I use with AccuPOL?
A: We recommend a final concentration of 0.2 mM of each dNTP.
Q: What is the error rate and speed of AccuPOL?
A: The error rate is 1.5 x 10-6 errors/bp. The speed is 90-120 sec per kb.
AMV One Tube Hot Start RT-PCR Kit
Q: Can I use random hexamers instead of gene specific primers in the One Tube RT-PCR Kit?
A: No. You may be able to use random hexamers along with gene specific primers, but the kit wont work with random hexamers alone. The kit functions by using one of the two gene specific primers to perform a reverse transcription of one particular transcript, and then this specific transcript is amplified using the same primer as in the RT step, plus the second primer.
BL21 (DE3) Competent Cells
Q: Are BL21 (DE3) competent cells resistant to any antibiotics?
A: The BL21 (DE3) strain is sensitive to all common antibiotics.
Q: Can you clone in BL21 strains?
A: Well, you can -- success depends on how efficient the cloning is. If you are using a cloning method with high yield and low background, then you can transform straight in BL21 to save time. If you are cutting and pasting and screening, then you are better off using a cloning strain like GC5 until you get what you want, and then putting the constructed plasmid into BL21.
Q: What is the difference between BL21 and BL21 (DE3)?
A: Often when people say BL21 they actually mean BL21 (DE3). If theyre talking about T7 expression systems, you know its BL21 (DE3). The major difference between the two is that BL21 does NOT does contain the T7 RNA polymerase gene, so it cant be used with T7-based systems. BL21 will express proteins from trc, tac, lambdaPL, and araD promoters.
Q: Im a little worried about the toxicity of the proteins I want to express. Can I still use BL21 (DE3)?
A: For expression of toxic proteins, we recommend BL21 (DE3) pLysS competent cells. If youre uncertain whether the proteins are toxic, try BL21 (DE3) first. You get a higher level of expression with BL21 (DE3); there is a slight inhibition of induced expression with BL21 (DE3) pLysS when compared to BL21 (DE3).
Q: Are the BL21 (DE3) cells salt inducible?
A: No, you cant use salt to control protein expression with Ampliqon BL21 (DE3) competent cells.
BL21 (DE3) pLysS Competent Cells
Q: Are BL21(DE3)pLysS Competent Cells resistant to any antibiotics?
A: The BL21(DE3)pLysS strain is resistant to chloramphenicol.
Q: What is pLysS?
A: pLysS is a plasmid that has the T7 lysozyme gene (lysS) on it. The T7 lysozyme protein has two effects: it degrades the cell wall and it inhibits T7 RNA polymerase. Because T7 lysozyme inhibits T7 RNA polymerase, it helps to keep expression of the T7 promoter off until the IPTG is added, allowing greater control of expression.
Q: When should I use BL21(DE3)pLysS competent cells instead of BL21(DE3)?
A: BL21(DE3)pLysS allows tight control of background T7 expression. So if youre concerned about toxicity, BL21(DE3)pLysS is the better choice.
Q: Im not using a T7 promoter for protein expression. Can I still use BL21(DE3)pLysS?
A: No. You can still use BL21(DE3), but you should not use BL21(DE3)pLysS.
Q: After the heat shock step, you dont mention that the cells + DNA should be placed on ice for 2 minutes. Isnt this the standard procedure?
A: Ampliqon tested this extensively and determined that the two minutes on ice step following heat shock is unnecessary. It has no effect on the efficiency of the transformation.
Buffer Optimization Kit
Q: Should I change my cycling conditions depending on which reaction buffer I use?
A: No. We recommend that you use your standard reaction set up and cycling conditions and try each buffer -- as the only variable -- in the same set of reactions. Once youve determined which buffer works best, you may need to optimize the reaction conditions for your primer-template system.
Q: Your standard buffer is offered in a "Mg-free" version. Can I get the 10X Ammonium Buffer without magnesium in the buffer?
A: "Magnesium-free" Ammonium Buffer is available through Customer Service.
cDNA Direct
Q: Can I use more than 5000 cells per extraction?
A: Absolutely! 6000 to 10,000 cells will work fine.
Q: What is the minimum number of cells I can use per extraction?
A: Weve seen good results with 5 cells per extraction. Although 1 or 2 cells per extraction has worked in some instances, we dont suggest using less than 5 cells per extraction.
Q: Do I need to add an RNase inhibitor to the RT-PCR reaction with my cell lysate?
A: RNase inhibitors are present in your cell lysate from earlier processing steps in the cDNA Direct From Cells procedure. Additional RNase inhibitor is not required.
Q: Can I use the cDNA Direct From Cells RT kit with bacterial cells?
A: We dont have data on this application. It is uncertain whether the freezing cycle is sufficient to lyse the bacterial cell wall. However, after you re-pellet your cells, you could try resuspending the cells in 1X PBS containing 1 mg/ml lysozyme + 0.05% Tween 20. Let the cells sit for 15 min at room temperature before adding Cell Lysis Buffer I and continue with the procedure.
dNTP Sets
Q: How much dATP, dCTP, dGTP and dTTP should I use in PCR?
A: The recommended concentration of EACH nucleotide in PCR is 0.2 mM. If you are starting with 100 mM stocks of each nucleotide, your dilution factor is 100/0.2 = 500X. Thus, if your PCR volume is 1 mL, you add 2 uL of EACH nucleotide dATP, dCTP, dGTP and dTTP. From a practical standpoint, if you are starting with individual nucleotides, the best thing to do is make a mix and then add the mix to the PCR.
Q: How do I make a 50 mM dNTP Mix from 100 mM stocks?
A: To prepare 200 uL of a 50 mM dNTP Mix (12.5 mM of each), combine 25 uL of EACH nucleotide dATP, dCTP, dGTP and dTTP [100 mM] with 100 uL water. Final volume is 200 uL. If a larger volume is required, the components can be scaled up directly.
Q: Can I make more DNA in my PCR reaction by adding a higher concentration of nucleotides?
A: Maybe, but the buildup of pyrophosphate and the inhibitory effect of too much nucleotide may offset your possible gains. The best way to scale up yield from PCR is to increase the volume of the reaction, not raise the nucleotide concentration.
GC10 High Efficiency Competent Cells (comparable to DH10B)
Q: How much of a ligation reaction should I use in my transformation?
A: We recommend that you dilute your ligation reaction 3-fold in TE buffer and use 1 ul of the dilution for a 50-ul transformation. Its important to dilute your ligation reaction because components of the ligation may interfere with the efficiency of the transformation.
Q: How much pUC19 DNA should I use in my control reaction?
A: Use 0.1 ng pUC19 per reaction. The transformation reaction is saturated at greater than 0.1 ng pUC19 per reaction.
Q: I have some pre-made agar plates. Can I add X-gal and IPTG to these plates for blue/white screening?
A: Yes. One method is to add 40 ul of a 40 mg/ml X-gal solution and 30 ul of 100 mM IPTG directly onto an 100 mm agar plate. Spread evenly using a sterile spreader. Let dry about 15 min and then store protected from light. In short, if blue/white screening for recombinants is desired, the plates should include 100 ug/ml ampicillin (or the appropriate antibiotic), 40 ug/ml X-gal and 1 mM IPTG.
Q: What is the formulation of SOC Medium?
A: 2% Tryptone, 0.5% Yeast Extract, 10 mM NaCl, 2.5 mM KCl, 5 mM MgCl2, 5 mM MgSO4, 0.4% glucose. Add MgCl2, MgSO4, and glucose (filter-sterilized stocks) AFTER the medium is autoclaved. Filter the complete medium through a 0.2 um filter unit. The final pH should be 7.0.
Q: When I pulled out my transformation plates from the incubator the next day, I had a lot of little colonies around big colonies. What are they?
A: These are satellite colonies. They are not transformants. Incubate your plates for less time (18 hours is good, not more than 20), use more antibiotic, or use fresher plates to get rid of them.
Q: How stable are your competent cells?
A: If stored properly at -70 C, our competent cells are stable for at least six months to one year. Do NOT store competent cells in liquid nitrogen.
Q: After the heat shock step in the transformation, you dont say anything about incubating the samples on ice for 2 minutes. Did you forget this step?
A: We tested this extensively and 2 min on ice after heat shock simply has no effect on the efficiency of transformation. Were just trying to save you time wherever we can.
Q: What is the maximum plasmid size I can use with GC10 competent cells?
A: GC10 competent cells efficiently transform a wide range of plasmid sizes, from very small to quite large (up to 150 kb). Yu may use either GC5 or GC10 up to 20 kb, but for plasmids larger than 20 kb we specifically recommend GC10. For very large plasmids, we suggest that you use electrocompetent cells, like our Thunderbolt GC10 Cells, for the highest possible efficiency of transformation.
Q: Im doing TOPO TA cloning. Can I use GC5 or GC10 competent cells?
A: We recommend GC10 competent cells with TOPO vectors.
SuperPath GC10 ElectroCompetent Cells
Q: How stable are your competent cells?
A: If stored properly at -70 C, our electrocompetent cells are stable for at least six months. Do NOT store competent cells in liquid nitrogen.
Q: How critical is field strength for electroporation? Should I try different settings?
A: Field strength is important in electroporation. Field strength is usually expressed as kilovolts/centimeter, where kV is equal to the initial peak voltage and cm is equal to the size of the gap between the electrodes of the cuvette. Optimal field strengths vary based on the type of cell being electroporated. For bacteria, use field strengths of greater than 15 kV/cm. For yeast, use 6 to 8 kV/cm. For mammalian cells, use 0.5 to 2.5 kV/cm.
Q: Sometimes I get arcing when I electroporate bacterial cells. How can I prevent this?
A: There are several ways to prevent arcing. You want to avoid conductive ions; water is fairly conductive, so its best to keep the ratio of DNA to cells in your sample low (5% or less). For example, use 2 uL of DNA in 40 uL of cells. Also, make sure there are no air bubbles in your sample and avoid any condensation which may form on the outside of the cuvette.
Q: I have some 0.4 mm cuvettes. Can I use them to electroporate the SuperPath GC10s?
A: No, 0.4 mm is too wide. We recommend 0.1 mm cuvettes for a reason -- its the only way to achieve the proper field strength. Just as the heat shock is critical to chemical transformation, the pulse -- the electric field generated -- is critical to electroporation. Its got to be short and intense. We recommend a field strength of 19 kV per cm, which is achieved by a voltage of 1.9 kV and a 0.1 mm cuvette. Its impossible with most cell porators to achieve this field strength with a wider cuvette because the voltage is usually maxed out at 2.5 kV.
Q: When I pulled out my transformation plates from the incubator the next day, I had a lot of little colonies around big colonies. What are they?
A: These are satellite colonies. They are not transformants. Incubate your plates for less time (18 hours is good -- never more than 20), use more antibiotic, or use fresher plates to get rid of them.
GC5 High Efficiency Competent Cells (comparable to DH5alpha)
Q: I have some pre-made agar plates. Can I add X-gal and IPTG to these plates for blue/white screening?
A: Yes. One method is to add 40 ul of a 40 mg/ml X-gal solution and 30 ul of 100 mM IPTG directly onto an 100 mm agar plate. Spread evenly using a sterile spreader. Let dry about 15 min and then store protected from light. In short, if blue/white screening for recombinants is desired, the plates should include 100 ug/ml ampicillin (or the appropriate antibiotic), 40 ug/ml X-gal and 1 mM IPTG.
Q: How much of a ligation reaction should I use in my transformation?
A: We recommend that you dilute your ligation reaction 3-fold in TE buffer and use 1 ul of the dilution for a 50-ul transformation. Its important to dilute your ligation reaction because components of the ligation may interfere with the efficiency of the transformation.
Q: Im doing TOPO TA cloning. Can I use GC5 or GC10 competent cells?
A: We recommend GC10 competent cells with TOPO vectors.
Q: How do I calculate transformation efficiency?
A: Transformation efficiency is the number of colony forming units (cfu) generated by 1 ug of supercoiled plasmid DNA in a transformation reaction. A known quantity of pUC19 DNA is typically used as the control. Note that you cannot measure the transformation efficiency by transforming with a microgram; the transformation reaction is saturated at greater than 0.1 ng pUC19 per reaction. Transformation efficiency (cfu/ug) is calculated as follows: cfu on control plate / ng of control DNA plated X 1000 ng / ug. For example, 0.1 ng of control DNA (1 uL of 0.1 ng/ul, freshly diluted) is added to 100 ul of competent cells. 900 ul of SOC medium is added prior to expression. 100 ul (equivalent to 0.01 ng DNA) is then diluted in 900 ul SOC and 100 ul is plated (equivalent to 0.001 ng DNA). If 100 colonies are counted on the plate, calculate the transformation efficiency as follows: 100 cfu / 0.001 ng X 1000 ng/ug. The transformation efficiency is 1 x 10E8 cfu/ug.
Q: How much pUC19 DNA should I use in my control reaction?
A: Use 0.1 ng pUC19 per reaction. The transformation reaction is saturated at greater than 0.1 ng pUC19 per reaction.
Q: How stable are your competent cells?
A: If stored properly at -70 °C, our competent cells are stable for at least six months to one year. Do NOT store competent cells in liquid nitrogen.
Q: Im using the single-use GC5, 50 uL per transformation. Can I perform the reaction in the competent cells tube?
A: Absolutely. You still achieve high efficiency when transforming in the competent cells tube. But note that the HEAT SHOCK conditions should change slightly: 37 C for 30 seconds is optimal when transforming in the competent cells vial with 50 uL cells.
Q: After the heat shock step in the transformation, you dont say anything about incubating the samples on ice for 2 minutes. Did you forget this step?
A: We tested this extensively and 2 min on ice after heat shock simply has no effect on the efficiency of transformation. Were just trying to save you time wherever we can.
Q: When I pulled out my transformation plates from the incubator the next day, I had a lot of little colonies around big colonies. What are they?
A: These are satellite colonies. They are not transformants. Incubate your plates for less time (18 hours is good, not more than 20), use more antibiotic, or use fresher plates to get rid of them.
Q: What is the maximum plasmid size I can use with GC5 competent cells?
A: GC5 efficiently transforms plasmids up to 20 kb. If your plasmid is larger than 20 kb, we recommend that you use GC10 cells.
GC5 Value Efficiency Competent Cells (comparable to DH5alpha)
Q: How much of a ligation reaction should I use in my transformation?
A: We recommend that you dilute your ligation reaction 3-fold in TE buffer and use 1 ul of the dilution for a 50-ul transformation. Its important to dilute your ligation reaction because components of the ligation may interfere with the efficiency of the transformation.
Q: I have some pre-made agar plates. Can I add X-gal and IPTG to these plates for blue/white screening?
A: Yes. One method is to add 40 ul of a 40 mg/ml X-gal solution and 30 ul of 100 mM IPTG directly onto an 100 mm agar plate. Spread evenly using a sterile spreader. Let dry about 15 min and then store protected from light. In short, if blue/white screening for recombinants is desired, the plates should include 100 ug/ml ampicillin (or the appropriate antibiotic), 40 ug/ml X-gal and 1 mM IPTG.
Q: How much pUC19 DNA should I use in my control reaction?
A: Use 0.1 ng pUC19 per reaction. The transformation reaction is saturated at greater than 0.1 ng pUC19 per reaction.
Q: How stable are your competent cells?
A: If stored properly at -70 C, our competent cells are stable for at least six months to one year. Do NOT store competent cells in liquid nitrogen.
Q: After the heat shock step in the transformation, you dont say anything about incubating the samples on ice for 2 minutes. Did you forget this step?
A: We tested this extensively and 2 min on ice after heat shock simply has no effect on the efficiency of transformation. Were just trying to save you time wherever we can.
Q: Value Efficiency GC5 cells achieve an efficiency of > 10^8 transformants/ug. Do you offer them at a lower efficiency?
A: 10^8 transformants/ug is the lowest efficiency we offer for GC5 competent cells, unless you purchase them in the 96-well format. However, the price for Value Efficiency GC5 is comparable to other lower efficiency competent cells. You get better efficiency without paying extra for it.
Q: What is the formulation of SOC Medium?
A: 2% Tryptone, 0.5% Yeast Extract, 10 mM NaCl, 2.5 mM KCl, 5 mM MgCl2, 5 mM MgSO4, 0.4% glucose. Add MgCl2, MgSO4, and glucose (filter-sterilized stocks) AFTER the medium is autoclaved. Filter the complete medium through a 0.2 µm filter unit. The final pH should be 7.0.
Rapid Ligation Kit
Q: What is the size limitation of DNA that can be ligated?
A: No size limitation. However, lower efficiencies are expected with large vectors/inserts. Electroporation is recommended (but not required) for large constructs (over 20 kb).
Q: How much DNA is needed for a successful ligation?
A: 100 ng - 1 ug total (DNA plus Vector). We recommend that you start with 50 ng of vector, then add a 3-fold molar excess of insert for a cohesive-end ligation or a 5-fold molar excess of insert for a blunt-ended ligation.
Q: Can less DNA be used?
A: Yes, but a smaller reaction volume may be necessary (less than standard 20 ul). In this case, it will be necessary to use the same amount of enzyme as in the typical 20-ul reaction.
Q: Is the ligase in your ligation kit the same T4 DNA Ligase you sell as a stand-alone product?
A: Theyre not the same. They have different storage buffers, for one thing. The composition of the enzyme is different as well. The ligase included in our ligation kit is called "T4 DNA Rapid Ligase" to help distinguish the enzymes.
Q: Your literature states that the Rapid Ligation Kit is specifically formulated for transformation. What is special about the formulation?
A: Components of ligation reactions often inhibit transformation, i.e., the ligation works great but the efficiency of the subsequent transformation reaction is poor. So, weve optimized our ligation buffer for BOTH the ligation and transformation reactions. The specific formulation is proprietary.
Q: How much of the ligation reaction should I use in my transformation?
A: Use 1/10 volume of the ligation mixture in the transformation. Our standard ligation volume is 20 uL, so we recommend 2 uL of the ligation mixture (~ 5 ng) in 50 uL competent cells.
Q: Is the improved transformation efficiency with your ligation buffer only observed when using Ampliqon competent cells?
A: No, it is not necesssary to use the Ampliqon ligation kit in tandem with Ampliqon competent cells to get good results.
Q: Can I leave the ligation reaction at room temperature overnight?
A: Your ligation reactions should NOT sit at room temperature overnight. It will lower the efficiency of the transformation. If you want to store your reactions overnight, place them at -20 °C.
Q: Can my ligations sit at room temperature for longer than five minutes? How critical is the time of ligation?
A: Your ligation reactions are fine at room temperature for up to one hour. After that time, the reactions should be placed at -20 C. Longer than one hour at room temperature does not affect the efficiency of ligation [its 100% complete] but does lower the subsequent efficiency of transformation. For best results, proceed immediately with the transformation following the ligation.
Taq with Ammonium Buffer
Q: Why should I use an ammonium reaction buffer instead of the standard potassium reaction buffer?
A: Potassium reaction buffer was the original reaction buffer used with Taq DNA Polymerase. Taq tolerates potassium cations, but in many primer-template systems, particularily with GC-rich templates, you get superior amplification with an ammonium reaction buffer.
Q: Im using a genomic DNA template, amplifying under standard conditions. Im not getting any product.
A: How much magnesium (Mg) is in the reaction? With genomic DNA, you usually need higher levels of Mg. Try increasing the Mg in the reaction to 3.0, 3.5, 4.0 and 4.5 mM. Ampliqon Ammonium Buffer is offered with various magnesium concentrations for convenience, or use the 25 mM MgCl2 solution provided with the enzyme.
Q: Im seeing extra bands on my gel.
A: There could be several reasons why you are getting extra bands with your amplified product, but here are three easy things to consider: (1) Check how much magnesium is in the reaction. If you are over 1.5 mM, cut back to 1.5 mM. (2) Check the reaction after fewer cycles: 15, 20, 25. (3) Try about 30% less Taq in the reaction.
Q: Im a little confused about the labeling on the reaction buffer tube. It says something about 1.5 mM MgCl2, but doesnt the buffer have 15 mM MgCl2 in it?
A: The 10X concentration of MgCl2 is 15 mM -- thats what is in the tube. When you use the buffer at a 1X concentration, the final MgCl2 concentration is 1.5 mM.
Taq with Magnesium Free Buffer
Q: Does the Magnesium Free Buffer contain Triton?
A: Yes, 1% Triton X-100 is present in the 10X Magnesium Free Buffer. The only difference between the standard buffer and the magnesium free buffer is that the latter contains no magnesium.
Q: Can I purchase just the Magnesium Free Buffer, without the enzyme?
A: Yes, the Magnesium Free Buffer is available separately. The cat. no. is 62-6086-08. You get 4.5 mL of buffer [3 vials, each containing 1.5 mL buffer].
Taq DNA Pol 1.1X Master Mix Kit
Q: How much Taq is in 1.1X TaqComplete?
A: There are approximately 5 units of Taq per 50-ul reaction. So, the 100 reaction size contains 500 units of Taq. The 500 reaction size contains 2500 units of Taq.
Q: Can I use a smaller reaction volume [less than 50 µl]?
A: Yes. We also recommend using 18 ul of the 1.1X master mix in a 20 µl reaction.
TEMPase
Q: Does it employ an antibody?
A: No, TEMPase is a chemically modified Taq which will render it inactive at room temp.
Q: Is the heat activation step necessary?
A: Yes, heat activation at 95C for 15 minutes is needed prior to cycling to activate the enzyme. IT WILL NOT WORK UNLESS YOU HEAT ACTIVATE.
UniPOL
Q: What extension temp should I use?
A: For a three-step cycling procedure, we recommend that extension is performed at 72 degrees C. If you are following a two-step cycling procedure, we recommend 68 degrees C for 25-40 cycles, with final elongation at 72 degrees °C.
Q: Will UniPOL leave an A overhang for TA cloning?
A: Yes it will! You get a mixture of ends, but the overwhelming majority of the PCR products generated using UniPOL have dA overhangs at the 3-end, like Taq. UniPOL is an ideal choice for cloning when higher fidelity is needed.
Q: How much enzyme should be used per reaction?
A: This requirement will depend on your template. We recommend starting with 0.5 µL of UniPOL per 50 µL reaction.
Q: UniPOL includes two reaction buffers, Buffer A and Buffer B. Whats the difference between the buffers? Which buffer should I use?
A: The main difference is that Buffer B contains a stabilizer. In general, we recommend Buffer A for targets up to 10 Kb and Buffer B for targets greater than 10 Kb. For really long targets, the stabilizer may increase the thermostability of the polymerases, giving higher yields. Of course, the best option is to try both, to see what works better in your system.
Q: Should I include DMSO in my reaction?
A: You dont have to add DMSO, but DMSO may increase yields when amplifying extra long targets or GC-rich regions. The DMSO helps to protect DNA from depurination. The DMSO concentration should be titrated (ranging from 1 to 5%) since its effectiveness is dependent on the length, complexity and GC content of the target.
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