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Closing a Plasmid by Ligation

Closing a Plasmid by Ligation


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I have been having trouble with what should be a fairly simple ligation. I had a plasmid that I needed to cut part out of, so I designed a couple PCR primers to do that. The plasmid already had a BamHI site on 1 side of the region I wanted to remove, so I made the other PCR primer with a BamHI site on its 5' end with 3 extra Cs so the restriction enzyme would bind better. Here is a crude diagram of what I tried to do:

So I did the PCR and BamHI digest, it looked good on gel, so I purified it from the gel, got the concentration on the Nanodrop, and set up a ligation.

However, I keep getting zero colonies. I have tried a 24 hour reaction at 4 degrees, and 10 minute reaction at room temp. I am currently trying a reaction where I supplement the buffer with fresh ATP, because the buffers are a little old. If that works, problem solved, if not, I am asking how to troubleshoot this.

This seems like it should be relatively simple, because I have no insert, just trying to close an empty vector, which seems to happen on its own when I don't want it to and dephosphorylate the vector.

UPDATE:

The reaction with extra ATP did not produce colonies. I will keep trying. One idea I had was to do the PCR and then try blunt end ligation, since the Pfu polymerase should make blunt ends. Also got a new sample of ligase.


Ligation, plasmid rescue problem! - PLasmid transformation after ligation problem (Jan/02/2008 )

Dear All
I have been having a problem since 2 months. I am ligating a 2.7 kb inser and a 3.4 kb insert into a 8 kb vector. Enzymes ends: KpnI, EcoRV and XhoI. I used negative control for ligation (where I only ligate my cut vector) and it showed no colonies. All plates of ligation reaction (after transformation) show colonies and seem great. However, when I do the miniprep, and run the gel to confirm the plasmid, I get different plasmid sizes all over. Sometimes Ig et just s a

3KB fragment sometimes I get lower but never 14Kb. Thus the bacteria is growing in Amp and it looks like ligation is working (although I have no positive controL) but I dont get my desired plasmid.

I think it could be recombination problem. That is the ligation is working but plasmid is recombining in bacterial genome and then some parts just leave the genomic DNA. So, I tried growing it at lower temperature (32 or 30 instead of 37) and I tried lower shaking speed. I used DH5a and sbtl2 strains yet same results.

Anyone has any suggestion?
Please reply.
Thank you

could you confirm what the insert digestion for this 3 way ligation?
Is it

Next how many colonies are you screening? And by the way what kind of check digest did you conduct?
I would suggest that you conduct colony PCR (using a PCR reaction that amplifies across the junction between inserts) to sceen. As you are now well aware screening by miniprep is a very time consuming process. Wth colony PCR and a multichannel pipette you can screen 72 colonies easily.

Just to confirm matters, are the DNA fragments clean? After digestion, were the DNA fragments gel purified?

As you have suggested, one possibility is the recombination problem. I would try SURE cells to overcome. Also can you describe your ligation mixture.

Thanks for dropping a reply. Well I don't know about SURE cells, are these teh expensive fancy ones? It is not sold by invitrogen, is it?

Well i did different ligation mixtures but always the vector was low compared to inser. like at least 3 times.. and for once it was in equimolar. Does this answer yoru question?

could you confirm what the insert digestion for this 3 way ligation?
Is it

Next how many colonies are you screening? And by the way what kind of check digest did you conduct?
I would suggest that you conduct colony PCR (using a PCR reaction that amplifies across the junction between inserts) to sceen. As you are now well aware screening by miniprep is a very time consuming process. Wth colony PCR and a multichannel pipette you can screen 72 colonies easily.

Just to confirm matters, are the DNA fragments clean? After digestion, were the DNA fragments gel purified?

I am sorry, I didn't see your reply earlier.
It is Kpn1-insert-EcoRV-insert-Xho1- Vector

200 colonies for now and you can never imagine how boring is that. I have done different ligations as well. The DNA is not so clean. I don't know why but whenever I gel extract anything using Qiagen, the 260/230 is too low (ethanol and salts). I tried washing mroe than once with PE yet the same. 260/280 is normal. The amount I am getting back from Qiagen columns is much less than what I used to (when I worked in another lab). I order a new kit yet the same is happening.

I am still having a problem if anyone has any suggestioN!
I am uploading an image of a gel I ran of the plasmid i miniprep.

Thanks for dropping a reply. Well I don't know about SURE cells, are these teh expensive fancy ones? It is not sold by invitrogen, is it?

Well i did different ligation mixtures but always the vector was low compared to inser. like at least 3 times.. and for once it was in equimolar. Does this answer yoru question?

Sure cells are available from stratagene.

For some difficult ligations, i would use 5 -10 x more insert (1:5 or 1:10 ratio of vector: insert) in the ligation.

100 ng of vector and insert in 1:5 ratio. This may help.

For gel extraction, try invitrogen. I used to like Qiagen but now have the same problem as you are facing, so switched to invitrogen.

Thanks for dropping a reply. Well I don't know about SURE cells, are these teh expensive fancy ones? It is not sold by invitrogen, is it?

Well i did different ligation mixtures but always the vector was low compared to inser. like at least 3 times.. and for once it was in equimolar. Does this answer yoru question?

Sure cells are available from stratagene.

For some difficult ligations, i would use 5 -10 x more insert (1:5 or 1:10 ratio of vector: insert) in the ligation.

100 ng of vector and insert in 1:5 ratio. This may help.

For gel extraction, try invitrogen. I used to like Qiagen but now have the same problem as you are facing, so switched to invitrogen.

Malik
Do you think it is recombination problem? I tried DH10B Max efficiency and Sbtl2 cells still getting same result.

I think recombination could be a factor. Even if you try cells which prevent recombination, they do not completely prevent recombination. From what you have described, like having many colonies but of varying sizes, sounds like recombination. I have actually had problems with recombination and from what you described, it seems like that. I have used XL-gold and SURE cells from strategene for this problem. Other cells types might work too. Try increasing the total DNA in ligation and also have ratio of 1: 5 or 1:10 and give it a go. I solved by using very high conc of DNA in ligation like vector 150ng and insert ratio 1:5 and it worked for me. But think of other likely problems as well. Like over digestion of the ends.


DNA Ligation

Ligation of DNA is a critical step in many modern molecular biology workflows. The sealing of nicks between adjacent residues of a single-strand break on a double-strand substrate and the joining of double-strand breaks are enzymatically catalyzed by DNA ligases. The formation of a phosphodiester bond between the 3' hydroxyl and 5' phosphate of adjacent DNA residues proceeds in three steps: Initially, the ligase is self-adenylated by reaction with free ATP. Next, the adenyl group is transferred to the 5'-phosphorylated end of the "donor" strand. Lastly, the formation of the phosphodiester bond proceeds after reaction of the adenylated donor end with the adjacent 3' hydroxyl acceptor and the release of AMP. In living organisms, DNA ligases are essential enzymes with critical roles in DNA replication and repair. In the lab, DNA ligation is performed for both cloning and non-cloning applications.

DNA Ligase Fidelity: When does it matter?

High fidelity polymerases are everywhere&mdashbut why would you need a high fidelity ligase? And what do we even mean by &ldquofidelity&rdquo when we&rsquore talking about ligation? In this webinar, NEB Scientist and ligase expert Greg Lohman discusses mismatch ligation by DNA ligases and the molecular diagnostics applications that depend on the use of high-fidelity DNA ligases like NEB&rsquos HiFi Taq DNA Ligase to detect single base differences in DNA.

DNA Ligation

Ligation, the process of joining DNA fragments with a DNA ligase, proceeds in three steps. Learn more about the function of ligation with our quick tutorial animation.

What molar ratios should I use for DNA Ligation?

The optimal reactant ratio is contingent upon the downstream application.

Why do I need to add PEG to my DNA ligation?

Polyethylene glycol (PEG) is an important reagent in ligation reactions, find out why.

What are the best conditions for DNA ligation?

Find out how the downstream application dictates the best reaction conditions for ligation.

Are some ligations more difficult than others?

Ligation of blunt ends and single-base overhangs require optimized reaction conditions.


Designing plasmid vectors

Nonviral gene therapy vectors are commonly based on recombinant bacterial plasmids or their derivatives. The plasmids are propagated in bacteria, so, in addition to their therapeutic cargo, they necessarily contain a bacterial replication origin and a selection marker, usually a gene conferring antibiotic resistance. Structural and maintenance plasmid stability in bacteria is required for the plasmid DNA production and can be achieved by carefully choosing a combination of the therapeutic DNA sequences, replication origin, selection marker, and bacterial strain. The use of appropriate promoters, other regulatory elements, and mammalian maintenance devices ensures that the therapeutic gene or genes are adequately expressed in target human cells. Optimal immune response to the plasmid vectors can be modulated via inclusion or exclusion of DNA sequences containing immunostimulatory CpG sequence motifs. DNA fragments facilitating construction of plasmid vectors should also be considered for inclusion in the design of plasmid vectors. Techniques relying on site-specific or homologous recombination are preferred for construction of large plasmids (>15 kb), while digestion of DNA by restriction enzymes with subsequent ligation of the resulting DNA fragments continues to be the mainstream approach for generation of small- and medium-size plasmids. Rapid selection of a desired recombinant plasmid against a background of other plasmids continues to be a challenge. In this chapter, the emphasis is placed on efficient and flexible versions of DNA cloning protocols using selection of recombinant plasmids by restriction endonucleases directly in the ligation mixture.


TA Cloning

Another approach, called TA cloning , creates complementary single-stranded overhangs between the insert and vector by exploiting a secondary enzymatic property of Taq polymerase. Taq DNA polymerase has terminal transferase activity, which means it adds a single deoxyadenosine (dA) to the 3′-ends of double-stranded DNA. This is not true for all thermostable DNA polymerases, but is true for Taq polymerase. This reaction does not depend on the sequence. Compatible TA cloning vectors with T overhangs are supplied by different biotechnology companies. Instead of purchasing a prepared vector, the T overhangs on the vector can be generated by first cutting the vector with a blunt end restriction enzyme, and then by mixing it with a terminal transferase enzyme and dideoxythymidine triphosphate (ddTTP), which is missing the 3′ hydroxyl group. The missing hydroxyl ensures that only one T is added to the vector ends. Alternatively, the vector can be cut with restriction enzymes that generate a single T overhang.

The TA cloning procedure begins by producing the insert in a PCR reaction using Taq polymerase, which adds a single A onto the ends of the PCR product ( Fig. 7.04 ). Next, the PCR products are mixed with a vector that has complementary 3′ deoxythymidine (T) overhang. DNA ligase is added to connect the vector and insert. The same TA cloning vector can be used to clone any segment of PCR amplified DNA, and does not require the researcher to cut and purify the complementary vector as is done for restriction enzyme cloning. This procedure is especially useful when convenient restriction sites are not available.

When Taq polymerase amplifies a piece of DNA during PCR, the terminal transferase activity of Taq adds an extra adenine at the 3′ end of the PCR product. The TA cloning vector was designed so that when linearized it has single 5′ thymidine overhangs at each end. The PCR product can be ligated into this vector without the need for special restriction enzyme sites.

Taq polymerase generates single 3′ A overhangs with its terminal transferase activity. These can be used for cloning PCR products into TA cloning vectors.

Another variation of TA cloning, TOPO-TA cloning improves the above procedure by removing the ligation step ( Fig. 7.05 ). The first step is the same, the insert is typically created by PCR with Taq DNA polymerase so it has a single A overhang on both ends. The vector also has a T overhang to be compatible, but the overhang is created with topoisomerase I , an enzyme that recognizes 5′ (C/T)CCTT 3′ DNA sequences. It cuts the double stranded DNA vector at this sequence, but does not release from the DNA. Instead, topoisomerase I becomes covalently bonded to the 3′ end of the vector. When the 5′ end of the insert nears the 3′ end of the vector, the covalent bond energy is transferred to connect the vector and insert. Then topoisomerase I releases from the DNA, leaving the insert and vector linked together.

Figure 7.05 . TOPO-TA Cloning

When Taq polymerase amplifies a piece of DNA during PCR, the terminal transferase activity of Taq adds an extra adenine at the 3′ end of the PCR product. The TOPO-TA cloning vector was designed so that when linearized, it has single 5′ thymidine overhangs at each end, which is covalently attached to the topoisomerase I enzyme. When the 5′ end of the insert nears the 3′ T overhang, the energy from the covalent bond transfers to connect the insert and vector.


DNA ligation products: selection guide

The selection of a DNA ligation kit requires consideration of several factors. During cloning projects it is helpful to assess whether the ligation involves cloning a long insert, whether rapid ligation would aid the overall workflow, and whether the type of ends being ligated are blunt, A-overhang (TA-cloning), or sticky (cohesive).

Depending on experimental conditions, transformation efficiency can sometimes be improved by limiting ligation reaction volume and thereby using a smaller volume of ligation mix to transform cells. High-throughput cloning studies may require streamlined ligation reaction protocol approaches to save time and minimize sample handling steps. We offer a range of DNA ligation kit options for each of these scenarios, so use the following guide to help choose the ligation kit that is best for your experiment.

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Ligation impossible: help with my cloning protocol?

hello all, i've become so frustrated with a simple bit of cloning i thought i would turn to you guys for some advice.

i'm trying to sticky-end ligate (NdeI/BamHI) a 700bp fragment into an expression vector however after 2 months and 6 attempts i'm yet to get a successful recombinant plasmid. here is my protocol:

NdeI/BamHI cut 700bp fragment from pBlueScript and gel purify - insert has been sequenced and both restriction sites are present. i have attempted this digestion as either a double digest or two single digests checking from plasmid linearisation between. following purification, the insert looks fine on a gel.

cut vector (as above, have tried both double digest and sequential single digests), alkaline phosphatase treat and heat kill enzymes at 80°C for 20 mins. both restriction enzymes are linearising the vector when cut separately however i can't check that vector is being cut by both.

overnight ligate at 16°C (insert:vector 5:1), heat shock transform into DH5a cells and select for apramycin R. ligase and competent cells have been working fine with all my other experiments so it's unlikely the problem lies here.

every time i get a couple of background colonies with no insert (plasmid isolated smaller than empty vector). the gene i'm working with is a bacterial transcription factor however as it's being put under the control of a thiostrepton-inducible promoter i think it's unlikely that the gene product is being transcribed and is toxic to e. coli.

i've been told about something called "plasmid instability" where transformation just isn't possible, however i want to rule out any other potential faults in method before this. if you can think of anything else i can try please let me know. thanks!


PCR vs. plasmid vector cloning. wtf?! help!

can anyone with an understanding of molecular biology please help me out here? i need to understand the difference between polymerase chain reaction and plasmid vector cloning. i know they're both techniques used to amplify a specific DNA sequence hell, i even carried out PCR in the lab this week. but i've lost sight of the bigger picture and the internets are doing nothing except confuse me more. why use plasmid vector cloning in e-coli to amplify a sequence when you can just do PCR?! i realise this is probably a very basic gap in my knowledge but any insights would be greatly appreciated, and i'll remember the person who clears this dilemma up for me when i'm a famous molecular biologist (right now i'm just an undergrad.) peace out internets.

When you're cloning a vector into E. coli, the goal is to get a strain that will express whatever gene or element you're inserting. Then you can freeze it down and always have a strain that expresses x gene, etc. The PCR is used as a TOOL to do this. You use PCR to amplify the DNA segment/gene you want inserted into the vector for the ligation reaction, you use it to confirm the gene inserted correctly into the vector then, after transformation, you use PCR to help confirm the intact vector in indeed in the new E. coli strain. PCR amplification alone will not amplify a genetic element in vivo. Is that a little more clear?

There are a few reasons why you might clone rather than just running a PCR.

Fragment size, for a large fragment you can get better amplification efficiency through cloning due to the length of time it takes to amplify a longer fragment with PCR while you can just grow up the E. coli to get multiple copies.

In a vector you can save the fragment in glycerol stock for later work.

Primer specificity, if you clone various fragments into E. coli you can amplify all of them with the same vector primers while to amplify various fragments with PCR you will need primers for each individual segment you would like to work with.

I am sure there are some other reasons I'm missing, but this should help you.

another reason: sequence fidelity. (if you keep amplifying with pcr, mutations will arise. bacteria have dna proofreading mechanisms)

Some of these answers are missing somethings. Vector cloning can be used to produce the actual protein product, but it can also be used to amplify the gene of interest. The big advantage of growing up a plasmid in E. coli is that E. coli has error repair mechanisms. PCR is a fast easy way to amplify genes, but even high fidelity polymerase (e.g. Pfu) can make errors, and every cycle of PCR propagates those errors.

If you need a lot of a gene, without mutations (so the protein product will still be functional), you best bet is to clone it into bacteria. If you want to rapidly make enough DNA do diagnostics, or if you need to get enough DNA to transform the bacteria, then you need to do PCR.

PCR is much quicker, taking just a couple hours usually, and can start with very little starting material (down to just one copy of your gene). Growing up a gene by transforming it into bacteria takes longer (at least a day), and is not very efficient, so you need a lot of the plasmid containing your gene. Usually, you need to do a limited number of rounds of PCR to get enough copies so that some will be taken up by your bacteria, but you want to minimize the PCR to avoid opportunity for errors.

These are good comments. I one important factor not mentioned is that It's relatively easy and inexpensive to insert plasmids into E. coli. Imagine that you are interested in a particular gene and a mutation associated with it. Using PCR and customized fragments you can create a copy of the regular gene, and a mutated version that is ready to be inserted into a plasmid. Once it's ready, you can transform the bacteria to greatly amplify your genes of interest. E.coli usually doubles every half hour. In addition to growing your gene exponentially, designed into your fragments and plasmids are controls that allow you to select for only the correct insertions. Usually an common antibiotic is used. Any bacteria that have the correct insertion will have a bacterial resistance gene allowing it to grow, but not incorrect versions with no resistance. Make a giant batch of broth, and grow overnight. Then do what's called a plasmid prep, which breaks open all of the bacteria, and collect a purified volume of your sample. Do a couple more control checks, then you can do all sorts of stuff. sorry for any errors in grammar or spelling.

I'm also an undergrad, so correct me if I'm wrong.

The main point of plasmid vector cloning is to get E.coli to express the amplified protein. You can try to crystallize it to try to study its structure, or just harvest it. PCR is a necessary step in vector cloning. You need to have restriction enzymes that can cut your plasmid and gene of interest at appropriate ends, you amplify these sequences with PCR and try to ligate them together. Dumpɾm in with some E.coli and you electrocute them (electroporation - I think it opens up the membrane) a bit and some of them will have integrated the plasmid. Now you might be thinking, "how do I know which E.coli have integrated the plasmid?" because you obviously don't want a mix of them. It's been accounted for with what are known as selection markers. The plasmid contains a gene for the resistance of an antibiotic, such as kanamycin (NPTII gene, I believe). So you plate the E.coli onto the "selective medium" - which is supposed to contain the antibiotic. The E.coli without the plasmid die and you're left with the ones with it. This is just one example of course. Some plasmids use reporter genes (e.g. lacz). Instead of killing the E.coli, in the presence of a substance called X-Gal, the colonies of E.coli without the vector turn blue and the others are white. You can pickɾm out individually with a pipette.

At the beginning, things were very hopeful for recombinant DNA technology. It's how things like insulin and factor VIII, a clotting factor for hemophiliacs can be harvested much more efficiently (I think they used to take it from cadavers and whatnot, and insulin was harvested from pig organs). Since we're capable of designing and inserting any piece of DNA within a certain length into the vector, we can design proteins (including enzymes) as medicine. The problem is that these sciences are still in its infant stages. It's very hard to design proteins that will work. Think about all the things that goes into making a drug: you have to look for a target receptor, understand it, and confirm that it's the receptor. You have to look for potential lead molecules or proteins that will interact with the receptor AND do the desired job. You have to think about administration - what's the dose like? how long will it stay in the patient's body? Are there similar receptors elsewhere that it can affect and produce side effects? How much does it cost to manufacture it and can people afford it? You've got to make a lot of variants and see which one performs best and patent them. This is why drugs are so goddamn expensive. The rewards can be great, but the risks are huge. Most drugs are failures. You can dump millions of dollars and years of research to come up with a dud. Even if your product works, now you've got to market it, convince doctors to get their patients to try it out, and beat out the competition.

With PCR, all you get is more and more DNA, which is also useful for sequencing and genotyping and a fuck tonne of things in general. I think it's one of, if not the most important tool that biology's come across. Look into wikipedia for more details.

Hope that helps. In summary: plasmid vector cloning is used to express genes into proteins, and PCR is for amplifying specific sequences of DNA.


Closing a Plasmid by Ligation - Biology

Introduction

Using restriction enzymes

DNA methylation & restriction enzymes

Modifying enzymes

Miscellaneous

Cloning Procedures

Non restriction cloning
1. Gibson assembly protocol

Recombination
1. Gateway

Miscellaneous Procedures

Modifying enzymes:
1. Klenow,
2. Mung Bean

BioSupplyNet (searching for scientific supplies.)

Oligos Synthesis (Sigma/Genosis, Eisenberg Bros, MBC, Rhenium)

Vendors for DNA synthesis
When wishing to start a project, one of the main ways to obtain your gene of interest is to purchase it from one of the following vendors:

Protocol Sites

  • Protocol on-line
  • Cold Sprong Harbor protocols
  • Biotechniques (compiled by the Protein Expression and Purification Facility of EMBL, Heidelberg)

This site is maintained by Dr. Nurit Doron . Your comments are most welcome.


Plasmid transfer

You are trying to construct a plasmid to be transfected (introduced) into mouse tissue culture cells in order to find out whether a specific protein localizes to the nucleus or the cytoplasm. You have three purified plasmids as starting materials. The objective is to combine pieces of these plasmids to make one purified plasmid suitable for the proposed experiment.
One plasmid contains a promoter/enhancer that is active in the mouse tissue culture cells and downstream of that promoter (in the same plasmid) are suitable sequences for initiating translation of an RNA transcript to produce several amino acids of a protein. These amino acids include the sequence MDYKDDDDK (single-letter code) at the very beginning (M is the initiator methionine). That stretch of amino acids is called an epitope tag (it is actually the FLAG tag) because it can be recognized by an antibody when it forms part of a protein. Following the sequence encoding the FLAG tag are a number of restriction enzyme sites (BamHI, XbaI and EcoRI). Do not worry about the restriction sites being so close to each other. In reality that can make it difficult to cut with two of these enzymes together but assume here that there is no problem for an enzyme to cut at a site very close to the end of a DNA molecule.
The second plasmid contains a full-length cDNA for the gene of interest (let's call it GLI-1). Conveniently, there is an NcoI site (CCATGG) right at the initiator codon (the ATG sequence within the NcoI site).
The third plasmid includes a region of a different gene that acts, when transcribed, as a signal for the transcript to be cleaved and polyadenylated (which is important for mRNA stability and translation). The signal works if RNA is transcribed in the direction from XbaI to EcoRI.
The desired product includes the promoter and FLAG epitope coding sequence from the first plasmid connected to the GLI-1 cDNA (such that the whole GLI-1 protein is translated following the FLAG epitope to make a "tagged" fusion protein) and to the XbaI-EcoRI polyadenylation signal segment. It is not important if there are a few extra amino acids between the FLAG tag and the rest of the normal GLI-1 protein and the success of the experiment does not depend on the nature or length of the 3' UTR (untranslated region) of the mRNA made in the mouse cells. You can assume that the restriction sites shown do not occur at any additional locations in the plasmids shown and that all of the plasmids have ampicillin-resistance genes. You can use any additional materials (oligos, enzymes etc.) you think appropriate. BamHI cuts between the Gs and NcoI between the Cs in their recognition sequence, on each strand.

(i) Describe how you would make the desired plasmid (WITHOUT using PCR)
In your answer be sure to make clear
(a) the exact sequences of any additional DNAs you use AND the exact sequence in your final construct between the FLAG epitope and the start of the GLI-1 cDNA.
(b) steps in the procedure including purifications (explaining why they are necessary or optional)
(c) how you will identify a correct product and
(d) how you will make sure that the correct product is indeed definitely correct (pointing out the most likely imperfection you may come across).

(ii) You have exactly the same objective as in (i) except that now
(a) you are allowed to use PCR
(b) the first plasmid has the same promoter/enhancer but does not have FLAG epitope sequence instead there is just a BamHI site followed by an XbaI site and an EcoRI site at that position (just GGATCCATCTAGATCGAATTC from the sequence shown in part (i)) and
(c) the second and third plasmids no longer have any of the indicated restriction sites (say because the NcoI site is now TCATGG, the XbaI sites are TCGAGA and the EcoRI site GAGTTC, though the precise reason is not important).

How do you make the desired plasmid that will encode GLI-1 with an N-terminal FLAG epitope?
Pay attention to the same four issues as described in (i), including specifying the length of any primers used and the location of their binding sites (sites of hybridization) where actual sequences are not known (include sequence in places where it is known).

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Attachments

Solution Preview

Please refer to the attached file.

(i) Double digest plasmid 3 (P3) with EcoRI and XbaI and run on the gel and cut out the 0.8kb band.
Double digest plasmid 1 (P1) with EcoRI and XbaI and run on the gel the intervening region will be lost as it is too small to visualize on the gel. That is ok. For size comparison, you can always do an EcoRI and XbaI digests separately. The double digest should be smaller than the single digests. Then, cut out the linear DNA that is double digested and ligate the P1 with 0.8kb fragment using the T4 DNA ligase. Run on the agarose gel and confirm the size, it should now be 0.8kb more than the double digest which is,

4kb. So, the ligated plasmid should be 4.8kb. Let us refer this as P4.
The DNA should be eluted from the agarose plugs, phenol chloroform purified to get rid of the salts and precipitated using 1/10th of 3M sodium acetate and 2.5 times 100% ethanol. Ligation should be performed with purified DNA only for good efficiency.
Then digest P2 with NcoI and XbaI and run on the gel. You will get a 2.2kb fragment beside other fragments of 3.0kb and 0.5kb. The 2.2kb fragment has NcoI and XbaI sticky ends. Now design a linker that has the BamHI site (GGATCC) and three nucleotides (ttt) that will abolish the NcoI site but will not put the coding region out of frame and retains the ATG .

Solution Summary

Plasmid to be transfected into mouse tissue culture is discussed in the solution.