JoVE Biology

Homemade Site Directed Mutagenesis of Whole Plasmids

1, 2

1Department of Biology, Johannes Gutenberg-University Mainz, Germany, 2Proteomics division, AlPlanta, Neustadt an der Weinstrasse, Germany

    Downloads Comments Metrics Publish with JoVE

    You must be subscribed to JoVE to access this content.

    Enter your email to receive a free trial:


    Enter your email below to get your free 10 minute trial to JoVE!

    By clicking "Submit", you agree to our policies.

    Admit it, you like to watch.



    Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid. Here we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid using standard reagents.

    Date Published: 5/11/2009, Issue 27; doi: 10.3791/1135

    Cite this Article

    Laible, M., Boonrod, K. Homemade Site Directed Mutagenesis of Whole Plasmids. J. Vis. Exp. (27), e1135, doi:10.3791/1135 (2009).


    Site directed mutagenesis of whole plasmids is a simple way to create slightly different variations of an original plasmid With this method the cloned target gene can be altered by substitution, deletion or insertion of a few bases directly into a plasmid. It works by simply amplifying the whole plasmid, in a non PCR-based thermocycling reaction. During the reaction mutagenic primers, carrying the desired mutation, are integrated into the newly synthesized plasmid. In this video tutorial we demonstrate an easy and cost effective way to introduce base substitutions into a plasmid. The protocol works with standard reagents and is independent from commercial kits, which often are very expensive. Applying this protocol can reduce the total cost of a reaction to an eighth of what it costs using some of the commercial kits. In this video we also comment on critical steps during the process and give detailed instructions on how to design the mutagenic primers.


    Principle of Method:

    The site directed mutagenesis of whole plasmids explained in this video is a mutagenesis method which allows you to alter a cloned target gene by substitution, deletion or insertion of a few bases directly into a plasmid. It works by amplifying the whole plasmid, in a non PCR-based thermocycling reaction. During the reaction mutagenic primers, carrying the desired mutation in form of mismatches to the original plasmid, are integrated into the newly synthesized plasmid. After removal of the original plasmid from the reaction the mutated plasmid gets transformed into E. coli. The following steps are for screening purposes only, because the mutation efficiency of this method is not 100%. The whole procedure takes three days, but the main part can be done within one day.

    1.0 Day one:

    1.1 Thermocycling reaction:

    Original Plasmid: This site directed mutagenesis protocol works best with plasmids up to 10kb. Larger Plasmids are a bit difficult to mutate with this method and may take some patience and adjustment of the thermocycling conditions and/or competent cells Furthermore the plasmid which you work with must be isolated from a dam+ bacteria strain. For the thermocycling reaction you will need 10 to 60 ngs of the plasmid you want to mutate.

    Primers: Before you can set up your thermocycling reaction you have to have your primers at hand. You need about 150 ng of every mutagenic primer It is ok if you take 1.5 µl of a 1:10 diluted 100 pM stock. There are a few simple guidelines you must consider when designing your mutagenic primers:

    • The primers should be complementary to each other
    • The primers should be between 25 and 45 nucleotides in length
    • The mutations in form of mismatches to the original plasmid must be contained in both primers
    • The mismatches should be centered in the primer and flanked by at least 8 nucleotides on each side
    • The primers should have a GC-Content of at least 40%
    • The primers should end 5 prime and 3 prime with one or more Gs or Cs
    • The primers need not to be phosphorylated, nor do they have to be FPLC or PAGE purified, merely desalted they should be.
    • For calculation of Tm you don’t need any formula but Tm on the shipping certificate should be higher than 60°C.
    • For screening purposes it is practical to insert or delete a restriction site with your mutation. Because of the degeneration of the genetic code there are many possibilities to insert a restriction site with your desired mutation. The website of the New England Biolabs provides a tool to find such an appropriate restriction site. Simply enter your primer sequence coding for the amino acid sequence you desire, using the ambiguity code for DNA. The enzyme finder tool will tell you which restriction sites can be introduced parallel to your desired mutation. If you can’t find any possible or practical restriction site by this way, you may insert any new restriction site without concerning the genetic code at the site of mutation. Thereafter you can use this plasmid as template for a series of mutations in which this restriction site will be deleted by insertion of the new mutagenic primers. This approach may be very convenient if you’re planning to do a series of mutations at the same site of the plasmid.

    DNA-Polymerase: For this method you need a thermostable DNA-Polymerase which exhibits 3’–5’ Exonuclease activity and creates blunt ends We always use recombinant Pfu-Polymerase from Fermentas. If you’ve got problems with the amplification steps you may try a higher quality polymerase, but for most purposes the standard Pfu will be enough. Complete the reaction with dNTPs, polymerase buffer and water.

    1.2 Thermocycling conditions

    The thermocycler should be set up in the following way. The initial and recurring denaturation phase is set to 30sec at 95°C.

    The annealing temperature of the primers must not be calculated with complicated formulae. We usually set the annealing temperature to 55°C and the annealing time to one minute. For primers of 25 to 30 nucleotides which you will be using mostly, this works for us 95% of the time. Anyhow if this should not work, simply try varying the annealing temperature between 50- and 60°C.

    The elongation temperature depends on the polymerase you use. In this demonstration we use the Pfu-Polymerase from Fermentas, which calls for a elongation temperature of 72°C. The elongation time varies according to the size of the plasmid. We always calculate 1 min per kb, and add one extra minute to that time. For example, for a 9kb plasmid we would choose 10min as elongation time.

    18 cycles are sufficient to create enough mutated plasmid for the further use. Keeping the number of cycles low, also saves you time.

    Figure 1

    1.3 Gelcheck after Thermocycling reaction

    The successful amplification should be checked by performing an electrophoresis. Directly after the cycling has finished, load 5 µl of the reaction onto a 1% TAE agarose gel. If the amplification was successful, you should see a distinct band. However if the newly synthesized DNA is not clearly visible on the gel, you may try to precipitate the whole reaction and use it for transformation in the next step. For us this has seldom worked but it is worth to give a try. The best thing to do if the reaction does not work is to adjust the annealing temperature.

    1.4 DpnI digestion:

    Before transformation the original plasmid which served as a template must be removed from the reaction to prevent strong background. This is done by restriction digestion with DpnI. This restriction endonuclease cuts only methylated plasmid. Its recognition and restriction site is the sequence GATC whereas A has to be methylated. When digesting with DpnI only the original unmutated plasmid which was isolated from a dam+ strain gets cut, the newly synthesized mutated plasmid which is not methylated is not affected by DpnI. Simply add 1-2 µl of DpnI to the reaction and incubate it at least one hour at 37°. When using the Fast digest DpnI the incubation time can be decreased to about 15 minutes. The quality of your DpnI and the incubation time of this restriction digestion greatly determines how strong your background with unmutated plasmid will be.

    1.5 Transformation:

    After DpnI digestion the plasmid is ready for transformation into the competent E. coli cells. Simply add 5 µl from the DpnI digestion reaction into the competent cells and perform the transformation as recommended in your lab. In our case we use the heat shock procedure by incubating the bacteria-plasmid mixture on ice for 30 min and then heat shock it at 42°C for 90 sec. After adding 200 µl SOC-solution, the bacteria are incubated, vigorously shaking, at 37°C for 1 h. After 1 hour the bacteria are plated on selecting agar media.

    2.0 Day two

    2.1 Screening of clones part 1: Selection of clones

    Because the mutation efficiency isn’t 100% you need to screen for your mutants. We do this by restriction digestion in which we check for presence or absence of the restriction site which we added or deleted through primer integration. For this purpose we usually pick up eight colonies and grow them over night for plasmid preparation on the next day.

    3.0 Day three

    3.1 Screening of clones part 2: Restriction digestion with marker enzyme

    On the third day you perform a Mini Prep and subsequent restriction digestion with your marker enzyme. On the gel you then can see which one of your clones carry the desired mutation and which ones not.

    The screening method using restriction digestion works very well. Nevertheless, you should confirm your successful mutation by sequencing.

    Subscription Required. Please recommend JoVE to your librarian.


    Site directed mutagenesis is a mutagenesis-method which provides a fast way to mutate a gene carried by a plasmid. The whole reaction can be done in only one day. With this method, it will need only a pair of complimentary primers carrying the desired mutations and a proofreading polymerase such as Pfu-Polymerase. The newly synthesized plasmid can be separated from the parental plasmid by digesting the reaction with the restriction enzyme DpnI. This enzyme digests only the methylated DNA. Therefore only the newly synthesized plasmid can be transformed. In this tutorial we demonstrated performing a site directed mutagenesis by using a homemade mutagenesis kit. The benefit of this kit is the cost saving while the effectiveness remains. The critical points of this method are the primer design and the annealing temperature. In case the newly synthesized DNA can not be visualized after electrophoresis, which may indicate the failure of the method, one can try to precipitate the whole reaction and use it for transformation in bacteria. However the best way to solve this problem is trying to optimize the annealing temperature or increase the length of the constant region in the primers. Further, using a high quality polymerase or restriction enzyme may also improve the sensitivity of the reaction. The competent cells also are a key factor which effects the successfulness of this method. Therefore middle (107 cfu/µg DNA) or highly (109 cfu/µg DNA) competent cells are preferable.

    Although in this tutorial we did not demonstrate the deletion or insertion by using the homemade kit, we were successful to do these in our laboratory. We were able to successfully substitute, insert or delete at least 3 bases at the same time.

    Subscription Required. Please recommend JoVE to your librarian.


    The authors have nothing to disclose.


    Name Company Catalog Number Comments
    Pfu polymerase (recombinant or native) Fermentas EP0571 or EP0501
    dNTP mix 10 mM Fermentas R0191
    DpnI or DpnI Fast digest Fermentas ER1701
    primers Invitrogen desalted


    1. Papworth, C., Bauer, J. C., Braman, J., Wright, D. A. QuikChange site-directed mutagenesis. Strategies. 9, 3-4 (1996).



    I know your publication is not about a kit, but I was wondering if you can help me. Using a kit, I get through the amplification step with a product, but the transformation is unsuccessful. The control plasmid I use in every transformation (has not undergone amplification) gives positive results. I tried lengthening the extension time, yet still, no colonies. Do you have any idea what could be wrong? I've heard about duplication of primers during amplification. Do you think this could be the problem? Unfortunately, I don't have a restriction site where I need the mutation. Thank you for any help!

    Posted by: AnonymousJuly 23, 2009, 2:11 PM

    Another reason why the transformation is not working, could be the size of your plasmid. With large plasmids we also have problems in transformation. Try using very high competent cells or even electroporation. Hope I could help you.

    Posted by: AnonymousJuly 31, 2009, 8:04 AM

    Hi Leah, which kit are you using? DŒs it use the same principle as we use? There are other kits on the market where you have to perform ligation prior to transformation (Phusion kit from neb). You may also check if it actually is the right plasmid you are amplifiying by performing restriction digestion with it after amplification and comapre it to your input plasmid. Otherwise you could use your input plasmid as control plasmid in transformation to check if it gives colonies. To help you more, I need more information about the kit your using and any points were you vary from the original protocol.
    Best wishes

    Posted by: AnonymousJuly 24, 2009, 4:56 PM

    Mark, I was using a Stratagene Kit. I used my input plasmid as the transformation control, and that dŒs give colonies. I gave up on the kit and tried your method. I get a few colonies, but now am not sure if the plasmid was mutated properly. It's a long and complicated discussion! I would type it if you have time to read it! :)

    Posted by: AnonymousOctober 6, 2009, 11:59 AM

    Hi Leah, you can check your succesfull mutation by restriction digestion if you inserted or deleted the appropriate restriction sites with your mutagenic primers. If you are not able to introduce the sites together with your desired mutation you could first introduce a restriction site and then delete it by introducing your desired mutation. The introduction of a restriction site could then also serve as a control if the method is working. This would be the cheapest way. The easiest way would be to introduce your mutation without restriction marker and verify your successful mutation by sequencing of a number of colonies. When you choose this approach you could enhance the DpnI digestion to reduce background. However, you should always verify your mutation by sequencing before moving on with your experiments.
    Feel free to type your problems up, I would be happy if I could help you. Best wishes

    Posted by: AnonymousOctober 8, 2009, 3:40 PM

    Unfortunately, I can't insert a restriction site because the area I would like to mutagenize is in the replication region, in the antisense RNA region. I went ahead and tried again with different primers and over and over, I get no colonies after transformation. I use about 60ng DNA, I have varied the annealing temp, tried longer extension times. Maybe I'm not using enough dNTPs? I'm pretty terrible at some of the math regarding concentration. :) I have a 40mM stock that I've been taking .5uL and putting it in a 50uL reaction. Should I be using 1uL? I'm also wondering if maybe the added mutations cause so much of a difference in the RNA that it can't function and thus, cannot be established in a cell? I'm getting minimal help from my lab, so any insight would be very much appreciated.

    Posted by: AnonymousNovember 6, 2009, 2:59 PM

    Hi Leah, do I understand you right, your region of intrest lies in the origin of replication of the plasmid? If your region of intrest is not essential for plasmid maintainence you can try to mutate it and create a restriction site as a positive control. This one you can then mutate to your desired sequence, then using the absence of the restriction site as assay for succesful mutation. If your region of intrest is in the origin of replication and dŒs not allow any big changes you could verify your mutation by sequencing. Regarding the dNTPs, polymerases usually require a concentration of ²00µM or lower per dNTP. So just take 1µL of a 10mM dNTP mix for a 50µL reaction. Do you get a product on the gel after the reaction, check 5µL on a gel, you should see a distinct band. You may also try using electroporation for transformation, or try a different polymerase (proofreading, non strand displacing and producing blunt ends). Hope this helps you. Good luck.

    Posted by: AnonymousNovember 11, 2009, 1:04 PM

    I was wondering if you need to do ligation with this protocol.
    Also, if the primers are complementary, is there much likelihood of producing primer dimers, and how can we overcome that?

    Thanks for you help

    Posted by: Jyoti D.March 16, 2010, 12:01 AM

    ligation is not needed in this protocol. Normally no problems are observed with primer dimers if you stick to the protocol (ca ²5-45 bp primers with at least 8 perfectly matching bases on either side of the mismatched region). If you have problems, you can try to make the primers shorter. By the way, purification of primers (HPLC or PAGE) will also enhance the efficiency and accuracy of the reaction.

    Have fun

    Posted by: AnonymousMarch 16, 2010, 5:53 PM

    i ordered the finnzyme,s (phusion) kit for site directed mutagenesis. and in that kit the primer designing was different. i have primers which are designed back to back and they are 5' phosphorylated(for lagation purpose). kindly guide me if i can use these primers with this protocol. coz unfortunately i didn get that kit. i have to complete this part of my work in a week. i m relly worried.

    Posted by: AnonymousJune 2, 2010, 2:23 PM

    those primers are not overlapping and one of them have mutation (single). in taht kit they used hot start DNA polymerase. i have used the pfu polymerase. but got ambigous result. there were ² bands in pcr product. one of 3.5kb(size of vector(ptz) +my gene) and a 3 kb band. i wonder Y pcr produced ² bands :(

    Posted by: AnonymousJune 2, 2010, 2:31 PM

    Hi Aliya, I would assume that you can exchange the phusion polymerase for the standard pfu (and reverse)as they both have the same basic functions (generating blunt ends, no strand displacing activity, proofreading). Also the orientation and design of the primers can vary a lot between different mutagenesis protocols available, so there are definetely many ways to mutate a plasmid. In your case I would say it's ok to use pfu instead of phusion as long as you use the correct cycling conditions. If you get two bands, you could either play around with the cycling conditions and/or primer design or you could just simply extract the band of correct size and use it for the ligation. Hotstart polymerases have several advantages (mainly higher specificty) but they are not necessarily needed. If you still have problems, you may also order new primers and stick with the protocol above. Hope this will help you.
    Have fun

    Posted by: AnonymousJune 7, 2010, 4:48 PM

    Hi Mark, i have recently ordered new primers (overlapping primers having mutation in middle of the sequence) but i i can,t add a restriction site in them coz mutation is in middle of my gene. and in ur protocol u didn mentioned about the size i mean the plasmid plus insert which u r using. my plasmid is of ².8 and my gene is about .5kb. would this procedure work for this size of DNA. and how the nick protroduced in the strands would be sealed in the end. and another question is in fermentas prescribed manual for pcr wd pfu polymerase it is mention that the extension time should be ²min/kb. which makes almost 1² min for my DNA. what should i do now.
    thanx for ur guidence

    Posted by: AnonymousJune 15, 2010, 1:55 AM

    Hi Aliya
    If your desired mutation is in the middle of the gene and you dont want to insert "silent" mutations you can check by Sanger sequencing of a few clones. Mostly all of them carry the correct mutation. The size of your plasmid is totally fine for the protocol. For thermocycling you should definetely stick to the protocol of the supplier, if pfu is too slow for you just take phusion. Also I'm sure that not all 18 cylces we recommend in the protocol are needed. But generally the cycling times are quite long in this protocol, this is normal. The nick in the plasmid is repaired upon transformation in the bacteria.
    Have fun

    Posted by: AnonymousJune 15, 2010, 4:06 AM

    Hi Mark
    thanx a lot for ur guidence. i followed that protocol and i succeded to introduce my desired mutation in gene. i just did it in 1st attempt. i got positive results by sanger sequencing. thanx :)

    Posted by: AnonymousAugust 2, 2010, 1:42 AM

    can u tell me the mechanism of this method. how many copies of mutated plasmid will be produced by this method from pcr of single unmutated plasmid. i am a bit confused how the primers aneal and synthesis more copies from newly synthesized strands in next cycles of pcr.

    Posted by: AnonymousSeptember 29, 2010, 12:35 PM

    Hi Shazar,
    the plasmid amplification works by a non PCR based thermocycling reaction (no exponential amplification but a linear one, similar to sanger sequencing just without disrupting nucleotides). The amount of mutated plasmid will be increased by the amount of input plasmid with every cycle, assuming the reaction is 100% efficient. So if you use 50ng of template, after the first cycle there should be 50ng of mutated plasmid, after the second cycle 100, after the third 150ng and so on. The amplification only takes place on the original plasmid and not on the newly synthesized (to my best knowledge). The newly synthesized plasmid strands then form plasmids with nicks in each strand which lie at the 5 prime end of each of the primers. If you transform this nicked plasmid into E. coli the nicks get repaired and result in a normal plasmid. I think in the video there's a scheme of the amplification process. If you cant acces the video you can view the article which the viedo is based on (in german). It contains the same scheme.²6.lasso
    All the best

    Posted by: AnonymousSeptember 29, 2010, 5:28 PM

    I have the problem of primer dimer with this protocol and i did not find any band after gel then i redesigned the primer which are partially overlaping and it works well in the first time and i got a good band
    but when i repeat it again it failed -----i used the same primers but changed only one amino acid in the same position
    i do not know what should I do ?
    any advise will be helpful to me

    Posted by: AnonymousJuly 7, 2011, 10:43 PM

    Hello Friends..
    Well iam working on Site-directed mutagenesis of one gene coding for one enzyme,however i dont get desired mutant ,sequencing results in undesirable mutation beside desired mutation. I follow strategene quick change mutagenesis protocol. I use pET²8a vector.Could anyone please guide how to adress this issue..Thanks fr d help..

    Posted by: AnonymousAugust 12, 2011, 5:14 AM

    Hi Shad,
    have you made sure that the undesired mutation was caused by the mutagenesis and was not present already before? If the undesired mutation appears in a region outside the one you are mutating, simple check some more colonies. As the mutagenesis is based on a linear amplificaation undesired mutations should not be propagated and therefore not present in most of the colonies. However if the mutation lies in the region of your primers, go and check their sequence once again (on the tube). During mutagenesis your primers are incorporated into the new plasmid, so if these are corrupted you will never get a good result. You could as well go for HPLC purified primers, which diminishes the risk of having a mixture of primers. Hope this helps you.

    Posted by: AnonymousAugust 12, 2011, 8:24 AM

    Thanks dear for your kind reply,
    i have ensured the mutations were not present prior amplification. Mutations were not in the primer as it is FPLC- HPLC purified. I get many deletion and insertion kind of muations. i even changed the Polymerase, from Pfu polymerse to Ex taq polymerase, still didnt work out.. Please help me out with a nice solution to this problem.. always thanks..

    Posted by: AnonymousSeptember 6, 2011, 10:20 PM

    Ok, so the mutations seem to appear during mutagenesis and they are additional to the desired ones which you introduce with your primers, right? I'm not sure where the mutations lie relative to the primer but you must be aware that in sanger sequencing the read quality towards the end gets quite poor and might seem like a mutation. To ensure that what you are seeing is really an additional mutation, you could check the chromatogramm and also perform a second sequencing reaction from the other side to ensure you have high quality coverage off the putative mutation. But this only if the chromatogramm quality is poor. Otherwise, I would say, try a diffrent polymerase such as phusion hotstart. Maybe you could also go for a fresh aliquot of dNTPs. Sorry Shad for the delayed answer, but keep the comments coming if you've got problems. Hope this will help.

    Posted by: AnonymousSeptember 18, 2011, 8:43 AM

    Hey Thanks Dear Mark for responding to my queries.!
    Yea you r right, i get final additional mutations in the gene sequencing result[Macrogen]. The undesirable mutations found through out the gene not at any specific position relative to primers, when i checked by aligning the wild & mutant gene using BiEdit program. I change the polymerase to Ex taq polymerase, i get the desired band of expected size, however confirming its sequence results in failure due to same additional undesirable mutations.
    I dont have much idea about Chromatogram and about sequencing from other side? As we sent our sequencing to Macrogen Gene sequencing Comp. Could you please tell me little more about it.
    Always Thanks. Your suggestions surely helps me. Hope to keep getting your valuable guidance.

    Posted by: AnonymousSeptember 18, 2011, 11:10 PM

    Hi Shad, what I mean with sequencing from the other side is simply that you should make sure the sequencing reaction gives reliable results. So what we do when we check a plasmid is sequencing into the orf from both sides, 3 prime and 5 prime ends. You could as well use a primer which lies further into the gene but sequencing in the same direction. When you get your sequecing results there should always be file containing the unprocessed chromatogram for example .abi file. You can check by opening the file with program. Once you have the correct files, use these as input for the alignement and compare to the reads you get before mutagenesis. Extaq I don't know but generraly you should only use enzymes with proofreading activity (pfu based enzymes or kod should work). Ok and as well there is no expected band size because the reaction amplifies the whole plasmid. Maybe you can get some help from your labmates or the sequencing company regarding the quality of reads. Good luck and keep me posted. We will get it solved :-)

    Posted by: AnonymousSeptember 20, 2011, 3:55 PM

    I did this protocol for getting point mutants and i have been successful a couple of times. But recently i am noticing the sequence has multiple copies of mutated oligos. The primer sequence is concatenated and repeated several times in the mutated plasmid. The primers i used had a tm of 65 C and are ²8 bp long. What could be the problem ?

    Posted by: AnonymousOctober 7, 2011, 10:18 AM

    Hi Sankar, I have not yet encountered the problem you describe. However I could imagine it is caused by pairing of the primers at their ends. You could check if the sequences could allow such a pairing and the maybe add some more bases to dimish dimerisation. You could also try raising the annealing temperature. Keep me posted. Mark

    Posted by: AnonymousOctober 7, 2011, 11:55 AM

    Hi Sankar, I have not yet encountered the problem you describe. However I could imagine it is caused by pairing of the primers at their ends. You could check if the sequences could allow such a pairing and the maybe add some more bases to dimish dimerisation. You could also try raising the annealing temperature. Keep me posted. Mark

    Posted by: AnonymousOctober 7, 2011, 11:55 AM

    The problem I believe is, while the polymerase is amplifying, the primer spontaneously separates allowing the polymerase to copy the primer region on to a daughter strand. Since the DNA is circular two primer regions are close by in the daughter strand. The reverse primer could anneal in this region and the polymerase could amplify the daughter strand instead, creating multiple primer regions on subsequent PCR cycles. In a normal point mutation reaction, the amplified daughter strands should not act as a template.

    To solve this, i ran the cycle separately for the two primers in two different tubes and mixed the samples before dpnI digestion. I had several colonies and one in 10 were positive for the mutation. And sequencing gave no concatenated primer region in any of the positive clones.

    Posted by: AnonymousNovember 4, 2011, 5:33 AM

    Hey mutagenesis fans, if you have problems mutating large plasmids you can give this modified protocol a try. In a nutshell, the protocol works for well for large plasmids, makes use of KOD polymerase and only 6 amplification cycles are needed: (download quite slow but eventually works). Good luck and happy pipetting

    Posted by: AnonymousNovember 2, 2011, 2:17 PM

    I have amplified ss DNA library of 60 mer size by assymetric PCR method using gradient primer(F.Primer 100uM R.prmer-10uM). i could able to amplify ss DNA libray ,however fail to isolate amplified DNA band from the Gel slab by Crush & Soak protocol.


    Posted by: AnonymousJuly 16, 2012, 9:44 PM

    Hi all,
    I could't figure out how to us NEB tool to design primer bearing a new restriction site, which can be used for clones screening. could anyone share any tips on how to conveniently do so?
    Many thanks!

    Posted by: Oleksiy K.March 20, 2013, 1:38 AM

    Dear Oleksiy K.

    My name is Ana Egana and I am the Technical Support Manager at New England Biolabs. I would like to invite you to contact us directly at with your questions. We will be happy to review the tool with you and provide you guidelines on how to use it for your intended purpose. We look forward to hearing from you.



    Posted by: Ana E.March 26, 2013, 10:10 AM

    Your Figure 1 seems to be broken.

    Posted by: August P.March 21, 2014, 12:11 AM

    Post a Question / Comment / Request

    You must be signed in to post a comment. Please or create an account.


    simple hit counter