Restriction Digestion - Science topic

I would assume that the strong signal at 6kb is the vector and,as you say, is a strong signal compared to the insert because the longer vector traps more ethidium bromide so looks brighter. The question is why the vector is running at a higher size than expected. It might be worth a restriction digest of the strong band with an enzyme that cuts often and see what the size of the band is when its smaller restriction products are added together to see if this is just a gel running or secondary structure artifact

consider to have 100microgram/mL kanamycin antibiotics in your LB, then you may look back at transformation, like give it a try on ice for 30 mints, then for 45 sec at 42 degree Celsius in water bath, then 3 mints on ice and finally add one mL LB without kanamycin to the suspension, incubate at 37 degree Celsius at 220RPM for one hour and then plate for overnight. moreover consider your DNA is between 50-100ng/microliter. best of luck

Can you start with a different vector? Use TOPO-TA cloning? Use primers that add in a restriction site for a different enzyme for the insert? Site-directed mutagenesis to remove the restriction site in the vector? I do not understand why you must use this exact vector + this exact enzyme.

Something is going to have to be different since your current strategy simply will not work.

Talk with your supervisor, it's a waste of your time to insist you use a protocol that you know will fail.

Enzymes are cheap (mostly), your time is valuable. I'm sure you can come up with a reasonable solution.

Inheriting a plasmid project is basically a guarantee that something is going to be not as expected. yes, sequence it & you'll be able to solve your digest problems.

Good luck!

It also looks like you're loading too much in the well, 50 ng of plasmid should be enough to be seen with EtBr or SYBR Safe staining.

Most of these kits bind denatured dna on a membrane and then elute in water or dilute tris buffer. You can usually improve the yield of purified dna by heating the elution buffer to 70c and leave it on the membrane for twice as long as the recommended time quoted in the kit and if necessary do another elution to make sure that all of the plasmid has been released from the membrane

This how I proceed:

Vector reaction:

2.5 ug vector

2.5 ul Xho

5 ul 10x buffer

H20 up to 50 ul

Incubate 37C for 2 hours, then add 2.5 ul Nde (pipette up-down after addition) and continue with additional 1 hour.

heat inactivate and gel purify.

Insert Reaction:

1 ug Insert

1 ul Xho

5 ul 10x buffer

H20 up to 50 ul

Incubate 37C for 2 hours, then add 1 ul Nde (pipette up-down after addition) and continue with additional 1 hour.

heat inactivate and column purify.

Ligation

20-50 ng cut vector

1:1 to 1:10 insert

dNTP mix

H20 up to 19 ul

Add 1 ul ligase

(pipette up and down)

16 C 16 hour

Heat inactivate

Trafo 2 ul

That worked for me.

References:

Corless, E., Hao, Y., Jia, H., Kongsuphol, P., Tay, D. M., Ng, S. Y., & Sikes, H. D. (2022). Generation of Thermally Stable Affinity Pairs for Sensitive, Specific Immunoassays. In Yeast Surface Display (pp. 417-469). New York, NY: Springer US.

and attached file.

Yes, you can do it that way as long as you're putting the same volume of template into each sample. If the added template DNA volume is variable between samples due to concentration differences, then you need to make each reaction individually, because the volume of water added will change between each tube. But in each case, add the enzyme last.

There was an internal XhoI site in the vector which was creating issue. I got the clone by using different sites and another shuttle vector. Thanks everyone for their valuable suggestions.

Try precipitating your plasmid with LiCl and ethanol and use less water to resuspend it. Scaling up is for more ug of DNA

The PCR did not work (on the plasmid and on the colony), the expression also did not give any protein.

Potentially, the DNA fragment you cloned is not your target gene, but another DNA fragment you cloned.

If you want to confirm your result 100%, you would need to carry out sequencing.

You can mix and match chemicals/enzymes from different companies. Just be aware that enzymes from different companies may be different units/ul so be sure to check the tube so you are using the correct amount for your reactions.

Michael J. Benedik I have seen RE cut dna get bigger when the enzyme binds to the target dna but has not cut , Then it looks large ( runs slowly) but addition of 0.05%sds denatures the mixture and the dna runs more like its actual linear size

Thank u Ananthi Rajendran i will definitely try this.

add salt (NaCl or ammonium acetate (1/10volume of a 5M solution ph5) before adding 2 volumes of cold ethanol

Since I do not have a map of the construct I can not tell you which is which.

If the blue and white colonies are similar size then it should be ok, a few extra hours in the incubator would let them get bigger.

How big is your insert? Was the plasmid mini prep good so that you say plenty of vector size DNA?

Can anyone give me any suggestions for cloning a small fragment, in my case the 776 bp in pcdna3. 1(-). What should be the cloning strategy for this. How much vector and insert amount in Nanograms should be used

If your entire insert is challenging to amplify, you could check the 5' and 3' ends to verify those are in the correct vector. In brief, for the 5' end, use a F primer in the vector (and unique to that vector, not found in your pUC57) and a R in the 5' area of your gene. Use a similar strategy for the 3' end. Yes, it is possible to have 2 plasmids in 1 cell and get 2 bands (it's uncommon, but certainly can happen). What does your uncut DNA look like?

Michael J. Benedik Sir, When we try to precipitate the restriction digested products, will the salts not interfere the transfer of DNA onto the nylon membrane?

Among 20X SSC and 10X SSC, which is a better transfer buffer?

Thank you.

Finding colonies on a vector-only negative control plate in a cloning experiment can be puzzling and may indicate a few possible issues with your cloning procedure. Here are some potential reasons and troubleshooting tips:

To address these issues, you can:

By systematically addressing each of these potential issues, you can identify and correct the problem in your cloning procedure.

l Perhaps this protocol list can give us more information to help solve the problem.

I face the same challenge.

Given the low quantities of linearized DNA that will be present you definitely need some form of enrichment protocol. A miniprep kit might work.

Hi Swati,

As per my understanding the low expression of your protein could be attributed to various factors, including weak promoter strength, transcription or translation inhibition, poor solubility, proteolytic degradation, or incorrect folding. To address these issues, consider using a stronger promoter, optimizing expression conditions, employing a different expression system, incorporating protease inhibitors, and verifying protein folding. For protein purification, utilize a nickel-affinity column for His-tag-based separation, employ a cleavage enzyme for thioredoxin tag removal, and perform size exclusion chromatography to eliminate contaminants.

Thanks and Goodluck!

Shirish Kumar Gangber Thanks for posting the resolution of your problem. It could be helpful to someone in a similar situation!

Thanks a lot for your help.

, Amy Klocko ,Mohit Gupta

A picture would be very helpful. Have you tried each enzyme individually? That test might help parse out which enzyme is giving you the unexpected bands. BamHI is a bit notorious for off-site cutting. Sometimes a different buffer can help. I had this exact problem with a double digest of a pCAMBIA vector and the buffer change did fix the issue.

Try a PCR of the cDNA to see if your gene is being expressed.

Dear Lester Wu

if you refer to transformation in e.coli of a plasmid amplified by PCR (eg mutagenesis) supplemented with dpni to remove the background of template plasmid, you can directly transform it, dpni and its buffer do not interfere with transformation efficiency. If you are using genomic DNA as template for the PCR, the digestion with dpni is not necessary, since the genomic DNA, differently to the plasmid will not be able to enter and propagate into the E.coli cells.

How big a fragment does the digestions supposedly release? If this is a tiny fragment then you may just not be seeing it the fragment if you don't have enough DNA.

You might also confirm by PCR with the same primers whether or not your plasmid mini prep still shows the correct band by PCR.

1) If the sequence of protein B has repetitive elements, or ones that are homologous to other sequences elsewhere in the plasmid, then recombination within the plasmid can occur in the bacteria leading to the loss of the antibiotic resistance gene. This would make the transformed bacteria grow more slowly. If this is the issue, you can resolve it by using a recombination deficient strain of E coli (like NEB Stable), and by growing the bacteria at 32C instead of 37C.

2) Gibson assembly is a simple and very efficient cloning method that works well with a single restriction site, and it will always ensure correct orientation of the insert. NEB's manual on Gibson assembly is a good place to learn about it.

Heating does not remove supercoiling. It requires a nick or break in the DNA by an endonuclease or a topoisomerase or to remove it.

Before choosing digestion enzymes, it is better to first check the Nebcutter site and check the digestion sites of the enzymes. If you are sure of choosing the right enzymes, check the efficiency of the enzymes and their buffers on a different plasmid (if you are sure of the correctness of the extracted plasmid, their shearing or buffering enzymes will not work)

usually you can cut directly in pcr buffer but New England Biolabs website has a useful page showing the percentage expected cutting for many enzymes in pcr buffers and the ability of enzymes to cut close to an end of the amplimer (for cloning ) will also depend on how many additional bases there are after the cut site. Again NEB have a list of additonal bases needed for efficient cutting

Thank you Robert Adolf Brinzer sir for your valuable response.

What results were you hoping to see? I'm not seeing any labels to know what is was you digested & which (if any) samples are your controls.

You'll also want to include a DNA ladder.

Try again and good luck!

Have you checked the concentration and quality of uncut DNA before restriction digestion? Did it have sufficient concentration and its integrity was good? On gel, where you were checking digestion, did you use a DNA marker since it might help whether there is any issue during electrophoresis or not? If you cannot even see the marker on the gel, there must be something wrong with your gel or running buffer. If the DNA has good quality and concentration and there is no issue with the electrophoresis process you must get at least a band of backbone. I hope this will be helpful to you. Wish you the best!

Getting a pcr product only proves that some of the plasmids have your insert so it is possible that some plasmids exist without the insert and you were just detecting the positive clones, Running a gel with uncut, cut with one enzyme, cut with second enzyme and cut with both enzymes might give you an idea where the unexpected band is coming from but I think that Satyendra Mondal is probably right that it is one of the faster moving isoforms of the empty plasmid

Supercoiled dna will form a small sphere and run more quickly than linear dna which is long and has more solvation effects on its surface so linear will run slower in agarose gels

the rules for digestion are

not more than 0.2 microg/microl DNA (final)

not more than 10% volume of enzyme (over 10% glycerol you may have some unspecific cuts)

1u of enzyme /ug of DNA (with good enzymes now you can use 10x more if you want)

for 1h at 37°C (exept some enzyme cleave at 65°C ; to be checked)

use the buffer recommended by the provider

same for double digestion: check if the buffer of the two enzymes are compatible; if not start with the buffer with less salt then after inactivation of the first enzyme (10min 65°C generally) adjust the salt and volume and add the second enzyme

now with the high quality of restriction enzymes the rules are flexible (more enzyme, longer time (in that case the pb will be the quality of your DNA not your enzyme) and some provider have a "universal" buffer usable for a lot of enzymes ....

One way to approach this problem is to use a sequence alignment tool that allows for mismatches, such as EMBOSS Needle or Clustal Omega. These tools can align the two sequences and identify conserved regions where the recognition site for the restriction enzyme is located. Once you have the alignment, you can visually inspect the alignment to identify mismatches and assess whether they are due to insertions/deletions or lack of conservation in the recognition site.

Another approach is to use a tool that can search for motifs in sequences with mismatches, such as MEME Suite or FIMO. These tools can search for a specific recognition site in both sequences, allowing for mismatches and insertions/deletions. The output will indicate whether the recognition site is present in both sequences and if it is conserved.

In terms of determining a tolerance value, this will depend on the specific restriction enzyme and the level of conservation in the recognition site. You may need to experiment with different tolerance values to determine the best approach for your specific case.

These video playlists might be helpful to you:

Yes, it is possible to disrupt supercoiled pDNA after isolation from E.coli with restriction digest buffer. Restriction enzymes recognize specific sequences of DNA and can be used to cut DNA molecules at these sites. The restriction digest buffer contains the necessary components to ensure the activity of the restriction enzymes.

What is the host strain you isolated the plasmid from? If it is an EndA+ strain then your plasmid preps will have endonuclease contamination that gets activated when you add any restriction buffer containing Mg++

One effective way to circumvent this (other than using an EndA- strain) is to not treat your mini prep with RNase. The RNA will serve to inhibit the EndA activity.

It depends on the primers you used to determine if the target is present.

If they are internal to the gene then when you plate the bacteria and also the plasmid on agar if you take a negative colony for pcr then the wetting effect of the reagent mix will give you an amplimer so making the negative clone look positive. When you grow on the clone then the wetting effect is diluted so the pcr or digest will also be negative. For colony testing the best primer set is one primer on the plasmid and the other internal to your gene then only transformed colonies will give a pcr product

What does the no-ligase vs + ligase control look like? If you have same number of colonies without ligation, then you have background of uncut vector. If you only get colonies upon ligation then either you have some vector only singly digested and not double digested or else your insert is not so good so the frequency of ligation products is low. Did you screen a large number of colonies before saying you had no inserts?

Don't use water - pH should be slightly basic. The supplied elution buffer, 1x TE or 0.1x TE would work better.

When eluting, add 20ul of the elution buffer, transfer the columns to 70C heating block for 2-5 min, spin. Add another 20ul (without changing the collection tube), keep it at 70C for another 2-5 min, spin down. You can use lower elution volumes and do 3 elution repeats (I usually do no more than 2). This especially helps with larger plasmids.

Or you can evaporate some water from the eluted plasmid - speedvac, heat block, or vaccum chamber may help.

or maybe the sample was too much concentrated to allow enzymes to access the DNA and cut it.

try with lower concentrations, you could afterwards reconcentrate the sample by evapating some water.

for this new year, all the best

fred

hi, There can be many reasons, it depends on the how much DNA and enzyme you took, also for how long are you keeping the samples for digestion.

My process is a little different from yours. I don't extract the plasmid immediately after growing the colony, it's a waste of chemicals and time if it doesn't carry the gene. I do the PCR reaction first. If it has a target gene, I will extract the plasmid and take the next steps.

Good luck

Ask your professor for help with your homework. This site is for research projects, not cheating on assignments.

There are a few possibilities. It could be that the colonies you pick have both plasmids with an insert and plasmids without. So the PCR can detect the rare ones with an insert but the restriction digest is only showing no insert since most the plasmids don't have it.

It could also be that you have the right clone but your restriction digest is not working properly. Is the "plasmid" band you see (I presume you are only getting one band) the correct size for the parental plasmid, or could it be plasmid plus insert sized?

It might also be that you are getting false positives from the colony PCR if there was too much DNA in the transformation and enough was picked up from the plates when you picked the colony.

No, the standard free energy of hydrolysis of the phosphodiester bond in DNA is -5.3 kcal/mol. It requires energy to forge a phosphodiester bond, while to break one requires only enough energy to overcome the activation energy barrier, which is lowered by enzymatic- , acid- or base catalysis. Under physiological conditions, hydrolysis is further facilitated by the high water concentration.

Most vectors come with a "multi-cloning site" that includes "rare cutting" enzymes (ones that have >4 nucleotides as their recognition site). Choose some different enzymes and see if that helps.

It's more common than you would expect to have a cut site in your insert. If you know the sequence, check it again the restriction sites first.

Finally, unless you ordered the plasmid from a manufacturer, you shouldn't trust that you have been given the correct vector. Sequence and verify EVERYTHING. Better to be confident now than to realize in 2 years you've been using the wrong plasmid.

BL21 cells express EndA gene which degardes the DNA of choice. Try using other cells like DH5alpha. Chosing the right strain is very critical.

Hi Zachary Spaulding !

The PCR products could be either purified or gel extracted depending on your downstream work. As your starting material is a PCR product, you could pool the samples (say at least 5 to 6 replicates- around 100 or 120 ul) so that you can have sufficient concentration and quantity. Run the pooled samples at a low voltage and elute the fraction. Ensure that you do each step with proper care starting from gel solubilization until elution. For the final step, heat the elution buffer/sterile water up to 50 or say 55 degrees and add it to the center of the column (do not touch it as you can damage). Add around say 20 or a maximum 30 ul (to get more concentration, you can still decrease the volume to even like 15 ul). Incubate the elution buffer for 5 min and then spin down the column. You will get your eluted product at a high concentration.

Hope it works for you! All the best

1. Are you checking the PCR products on a gel to see if there's one prominent band? If not you might just be cloning in incomplete/off target products.

Smaller fragments stain less brightly on agarose gels and in general ligate in easier than bigger fragments so even a band that looks 0.25x the brightness of your intended product could potentially be significant.

2. Are you cleaning up the PCR product after it's generated?

Primer dimers/small aborted extension products/nucleotides can actually clone into TA/TOPO vectors.

3. pBluescript has a high copy number and your insert could potentially be toxic.

At high enough copy numbers even inserts that are normally non-toxic can become problematic and this will select for clones with indels, transposon insertions, etc.

4. Sometimes blue/white screening also gives false positive clones because the X-Gal isn't even across the plate or a clone grew a little too fast to accumulate much blue product, or the concentration is low. Putting them in the refrigerator for a day or two can help identify false positives. I've definitely had "white" clones that turned faintly greenish-blue after storage at 4°. This is more of a "background vector" problem though.

1. Is the restriction digest on the whole plasmid minipreps you sent for sequencing or did you digest a PCR generated fragment? I've found that colony PCR amplifies extracellular DNA left over from my transformed ligation and gives me false positives sometimes.

2. How far outside the inserted fragment are your sequencing primers? Like are the sequencing results generating a nice alignment all the way across where your insert should be (an intact multiple cloning multiple cloning site for example) or is the read just trailing off before it gets to exactly where the insert should be? Don't know how experienced you are with sequencing, but on a good day for me Sanger sequencing generates 700-800 bp of usable data, but often falls far short of that.

If it's neither of those my guess would be that your vector's sequence is not actually what you think it is. This is actually really common in my experience, most of the time plasmid sequences are provided based on what somebody thinks the sequence is without that being verified. In that case I would send the miniprepped backbone vector and some of your positive clones to https://www.plasmidsaurus.com/ which uses long-read sequencing to sequence the entire plasmid for very cheap. If shipping to the United States is too much of a hassle, maybe there's an equivalent company in Europe.

This question is too general, and it needs more details. You can have several problems: not working enzymes, degraded DNA, not good enough analytical method, not an enzyme target in your DNA and more...

As the others have said, with many enzymes it will still work ok, but perhaps at reduced efficiency.

Thank you @Pierre Beguin for your valuable comments.

Hi

You seem to have too much vector to insert according to your ratio amongst perhaps some other things that might be contributing. I would swap the ratio around to have more insert to vector.

You can try this if you like.

I usually would first do the restriction digest of the vector and insert as you've done. Often I do it for 2hrs.

Then I would keep the insert on ice while Al-phos treating (phosphatase) the vector for 1hr.

Both Vector and insert are then run on a gel, cut, gel purified and then vector and insert are incubated in a 3:1 ratio (insert to vector) for 2 min on ice (optional), then subsequently ligation buffer and ligase added. The reaction can be done for 1hr or overnight.

Good luck and hope it works for you.

Annemarie

If your plasmid is large and if the total OD260 gives 448ng/ul then the actual insert concentration is much less than expected by the ratio of the insert size to the plasmid plus insert size. Also run an uncut sample before doing the next digestion to check that some of your nucleic acid is not RNA contamination from the bacterial host

Congrats! So glad I could help! :)

Yes this should be fine as long as the HB101 you transformed does not carry any plasmid, they will simply now carry your recombinant plasmid. Note that HB101 is generally suitable for cloning with the downside of being endA+, however, it your intention is expression and protein purification I would use a different strain.

The big smear at the end of the lanes looks to me more like RNA that remained from your minipreps but can not be certain. Did you run an undigested control of your plasmid to see what the DNA looks like before the digestion? I would recommend doing that to be sure you have plenty of intact plasmid.

The easiest way to solve your yield problem would be to PCR out the DNA segment that you want to insert into pTA927 CFLAG using appropriately designed primers that would provide the required restriction sites, plus of course a high fidelity DNA polymerase. After performing the PCR, gel purify the fragment, procced with restriction enzyme digestion to create cohesive ends and ligate with digested pTA927 CFLAG. Once you get the recombinant clones, check by colony PCR for the presence of the insert and send a couple of positive clues for sequencing to make sure the no spurious mutation has occurred during amplification.

Hi Leran, happy New Year to you too!

What I meant was to:

1. Fuse the gene already present in the Addgene plasmid (Gene 1)with your gene of interest (Gene2) by overlapping PCR adding the suitable restriction sites. PCR1 (gene1) + PCR 2 (gene2) = PCR12.

2. Digest the original plasmid removing the gene 1 and leaving cohesive ends for inserting the PCR12. Instead of amplifying this whole piece by PCR.

Anyway, I think that growing the positive control in liquid medium is a good idea. If possible try also another E.coli strain or richer media (SOC, YEB...)

After NEB HiFi assembly, you confirm the presence of the plasmid of interest by PCR. Do the primers used for this purpose bind to part of the insert fragment? if so, this might give false positive results but I get the feeling your primers don't bind the insert at all.

I had a similar experience when cloning in E.coli a vector for protein expression in yeast. There was no growth after transformation in E.coli. What we did was transform directly the ligation into yeast and it worked and yeast colonies grew perfectly. I understand this is not possible for you but maybe it helps you somehow...

I wish you luck for your experiment!

Hi!

I obtained this protocol provided by a company that sells that kit.

Purify DNA from cell or tissue samples by a desired method or commercial DNA purification kit. Reagents needed but not supplied: Nuclease P1 3M Sodium Acetate, pH 5.2 1M Tris pH 7.5 Alkaline Phosphatase Zinc Chloride For digestion, 15 µg of DNA in 100µL DNA hydration buffer or DI water is required. Protocol volumes can be scaled. 1. Prepare working solution of Nuclease P1 at 5U/mL in 40mM sodium acetate. Keep on ice. Remove aliquot of alkaline phosphatase, 10U/mL, from -20˚C. Keep on ice. Thaw normalized DNA samples (15 µg/100 µL). 2. Denature the DNA at 95-100˚C for 10 min. Cool completely on ice 5 min. Centrifuge for 5 sec or tap any condensate down into tube. Add 50 µL 40 mM sodium acetate pH 5.0-5.4, 0.4 mM ZnCl2 . 3. Add 50 µL of 5U/mL Nuclease P1. Invert tube to mix. Centrifuge 5 seconds or tap any condensate down into tube. Incubate at 37˚C for 30 min. 4. Adjust pH to 7.5-8.0 by adding 20 µL 1M Tris pH 7.5 to tube. Add 15 µL of 10U/mL alkaline phosphatase. Invert to mix. Centrifuge 5 sec or tap any condensate down into tube. 5. Incubate at 37˚C for 30 min. Boil samples for 10 min at 95˚C to inactivate alkaline phosphatase. Place samples on ice. Aliquot samples, 2 µg/tube and store at ≤ -20˚C until assaying. Samples should be diluted ≥ 1:4 with the diluted Assay Buffer prior to running in the assay.

Hope this helps!

It is hard to say if you have a poor yield without knowing the relative sizes of the plasmid and the insert. For instance if your plasmid is 30kb and your insert 300bp then of your 1ug DNA total there is only about 1/100th of the total signal is insert. I would cut more plasmid and do not run the gel for very long ( 1/4 the time of your picture) then you will still get good separation of plasmid and insert but the insert will be more compact and clearer but do check what yield you can expect given the relative sizes and you may find that this is a very good result. remember also your total volume is 50ul and if you are only running a fraction of this on your gel then the total yield will be greater