Change pBI121 promoter - (Aug/10/2007 )

Hi all

I want to cut pBI121 promoter (CaMV 35) with HindIII and XbaI.So I have some question:

1- If the promoter will be cut , should I see two band on my gel? I mean that, in consider, the CaMV is 853 bp Can I see a band for it?
2- And is it different to cut the plasmid with which on first (HindIII or XbaI)?

Thanks

---

Yes, you can visualise a 853bp band on a 1% agarose. Although for better resolution, you should run the DNA on a 1.2% or even a 2%.


No, not really. Only if the two sites are very close together (<10bp) then you may want to cut XbaI first followed byb HindIII. In any other situation, I would do a double digest, and cut the DNA with both HindIII and XbaI at the same time, in the same digestion mix.

---

Thanks perneseblue

Could you please tell me how much enzyme, plasmid, H2O and Buffer do you add in a double digesting reaction?
And if it is possible send me your gel picture. I want to know what I should see in mine.


Thanks again

---

I follow the philosophy of ‘It is better to have too much then too little.’ So the recommendations that I make below will be on the wasteful side.

15 ul plasmid DNA (aim is to get 10ug of DNA)
10 ul NEB buffer 2
5 ul NEB BSA (100x)
1.5 ul XbaI (~10U)
1.5 ul HindIII (~10U)
67 ul ds water

Incubate overnight at 37 Celsius. Gel purify the above in a very large well, made by taping several slots on the comb together.

At the moment I am not in the lab and thus unable to provide you a picture. But what you should expect is two clear band (if you are cutting out an insert).

Oh and do remember something about XbaI, this restriction site can be dam methylated by overlapping the dam methylation sequence. So before you start, please look up the adjacent sequence to the XbaI and see it your XbaI site is dam methylatable.

---

Thanks a lot for your advises. As you notice I checked XbaI neighbor's sequence, fortunately there is not an dam methylated site
By the way I don't have BSA know. It is possible to cut without this ?
And I work with Fermentas enzymes and buffers (Tango) , It is necessary to change enzyme and buffer volume for each brand?

Thanks again

---

No. Nearly all restriction digest buffers (Roche, NEB, fermentas) will be at 10x concentrations. It is practically an international standard. So there is no need to change buffer volumes. Enzyme concentrations are also nearly about the same, so no need to change the enzyme concentration (if you are using directly from the stock)

THere is already BSA added into fermentas buffer. So no need to add BSA saperately

---

Appreciate your kindness, Thanks

---

This question to pernese blue
I hope you're using pBI121 plasmid for your research. If you dont mind can you give me suggestions about ligation using pBI121 like what concentration of vector and insert ratio is best employed by you, then about the incubation temp and duration.. And best way to increase the efficiency of E.coli DH5 alpha cells to transform such a big vector.. Hope to receive your reply at the earliest................

---

Unfortunately I don't actually on plant plasmids. However for plasmids this size, i find concentration rather then ratio to be more important. Try getting your DNA as concentrated when conducting the ligation reaction. As for ligation conditions, the rector to insert ratio I would use is 1:1, to 1:3. I don't actually vary the ratio from standard.

I would also conduct this ligation as an overnight ligation at 16 celsius, using 0.5ul of T4 ligase in 20ul ligation volumes.

As for transformation efficiency, the only thing I can suggest is to use electroporation rather then chemical transformation. And to use company ultra competent cells such as Invitrgen Oneshot Genehogs, rather then home made cells. Using these two, I don't experience any difficulties.

After purifying your pBI121 vector, you should consider electrolution to extract the DNA from the gel matrix. It is a slow method, but you lose less DNA by electrolution then column extraction.

---

The concentration of both the vector and insert were very good. That even I load 2ul of eluted samples after restriction digestion, I'm able to see them on gel. I set a ligation rxn in the ration of 1:5. when I just ran the ligation mix on the gel, I was able to see bands corresponding to insert size, dimers of insert size and this appears a log increase of insert size and finally my vector.
When I transformed it I was able to get Kan resistant colonies but they were just the vectors alone without carrying the insert.
I would like to know how a plasmid which is double digested with various enzymes can religate? Also, How to prevent insert insert ligation? What can I do further to increase the ligation efficiency? I'm really bugged up with this problem... Please do help.

---