You will want to do a partial digest.  There are two ways you can do this, one by limiting the amount of enzyme you add to your reaction, the other by limiting the amount of time the enzymes are allowed to cut.

I have done a lot of partial digests and the optimal protocol that I use varies the length of time that the enzyme is allowed to cut. I set up a master mix of plasmid (~5 ug), RE buffer and water to give a final volume of ~80 uL.  I also prepare 4 eppendorf tubes that contain gel loading buffer (GLB)(usually ~2 uL of a 10X stock).  Also make sure that you have something handy that you can use to rapidly freeze your samples, either dry ice, liquid nitrogen, or an isopropanol:water (50:50) mix that has been stored in the -80.  The key is that you want to be able to stop the reaction very quickly.  You then add the enzyme (Sma1 in your case) (~0.5ul) and at various time points, (30s, 2min, 5min, 10 min) take 16 uL out of the master mix, add to the tube with the GLB, and then freeze it.  For the last time point (~20 min), add the GLB directly to the rest of the master mix in the original tube.  You then run your samples out (I usually load them in reverse order and do this quickly because as soon as the samples start to thaw, the enzyme will start to cut again.  Run out the gel and you should see a gradient going from mostly uncut plasmid in the first tube to mostly completely digested in the last reaction.  You can either cut the linear fragment (the one that has been cut at either Sma1 site) directly out of the gel, or you can do a another partial digest, this time stopping the entire reaction at the optimal time.  After extracting the linear fragment, you can then do a second digest with Sal1, and then extract the correctfragment.

Partial digests sound complicated but they are really quite easy and becoming proficient at them can really help your cloning.

Good luck,

J

You will want to do a partial digest.  There are two ways you can do this, one by limiting the amount of enzyme you add to your reaction, the other by limiting the amount of time the enzymes are allowed to cut.

I have done a lot of partial digests and the optimal protocol that I use varies the length of time that the enzyme is allowed to cut. I set up a master mix of plasmid (~5 ug), RE buffer and water to give a final volume of ~80 uL.  I also prepare 4 eppendorf tubes that contain gel loading buffer (GLB)(usually ~2 uL of a 10X stock).  Also make sure that you have something handy that you can use to rapidly freeze your samples, either dry ice, liquid nitrogen, or an isopropanol:water (50:50) mix that has been stored in the -80.  The key is that you want to be able to stop the reaction very quickly.  You then add the enzyme (Sma1 in your case) (~0.5ul) and at various time points, (30s, 2min, 5min, 10 min) take 16 uL out of the master mix, add to the tube with the GLB, and then freeze it.  For the last time point (~20 min), add the GLB directly to the rest of the master mix in the original tube.  You then run your samples out (I usually load them in reverse order and do this quickly because as soon as the samples start to thaw, the enzyme will start to cut again.  Run out the gel and you should see a gradient going from mostly uncut plasmid in the first tube to mostly completely digested in the last reaction.  You can either cut the linear fragment (the one that has been cut at either Sma1 site) directly out of the gel, or you can do a another partial digest, this time stopping the entire reaction at the optimal time.  After extracting the linear fragment, you can then do a second digest with Sal1, and then extract the correctfragment.

Partial digests sound complicated but they are really quite easy and becoming proficient at them can really help your cloning.

Good luck,

J

I agree with Jonathan, Partial digests are feasible but here since it seems you have 2 SmaI site within the MCS sequence (i.e very close to each other), it will be impossible to differentiate which SmaI site is being cut... you'll end up with a mix population no matter what. Then you can fully cut with SalI. I think you'd save a lot of time, and money, by re-ordering another primer! 

Re-designing primer with  unique site is needed to save your time.

@ Jonathan D Finn:

Thank you so much for your suggestion. I have tried it in a bit different way. First I did digestion with Sal-I followed by reaction clean up, then digestion with Sma-I (@ 30 degree for 1 hr whereas recommended is 37 degree c. ). Post ligation and transformation I got 11 colonies on selection plate, all them turn positive in colony PCR. I have set up the digestion of purified clones and waiting for the results.

Definitely re-designing a primer is better option but I want to solve this problem as it is acquiring a new skills and understanding.

Hi Nikhil,

You didn't make it clear if it is necessary that all those sequences remain (e.g., to maintain an orf or other reason).  However, if so, the way to clone this is to do a partial digestion as others have suggested.  Redesign your primers to take advantage of InFusin (Clontech) or Gibson Assembly (NEB) strategy.  With proper primer design your fragment should only clone into the SalI and correct SmaI sites.  If you don't have access to these cloning reagents, then clone after partial digestion and screen by colony PCR using primers to the pIRES vector + insert, spaced closely enough together to easily distinguish which SmaI site was retained.  Good luck.

Hi Nikhil,

While I hope that this works for you, I would be pretty surprised.  While lowering the temp from 37 to 30 degrees will slow the enzyme down, usually the amount of enzyme added is far in excess of what is necessary, and from my experience doing partial digests it seems like the RE always cut way faster than expected.

I also agree with Phillippe, if the two sma1 sites are close together, then this sort of partial digest is very hard to do as you cannot distinguish your plasmid that has been cut once from that cut twice.

I would suggest ordering alternative primers with unique RE sites.

Let us know how it goes!

JDF

Other comments here have suggested partial digestion by Smai in order to allow cleavage at only one of the two SmaI sites.  That can work.  But the reaction is difficult to control.  Below is an alternative suggestion which exploits the position of the SalI site relative to the two SmaI sites.

I assume that this question pertains to the "pIRES MCS B" shown at:

http://www.clontech.com/US/Products/Fluorescent_Proteins_and_Reporters/Coexpression/ibcGetAttachment.jsp?cItemId=17708&fileId=5877972&sitex=10020:22372:US

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Topic #1:  Contextual information regarding choice of the pair of restriction enzymes: SmaI and SalI:

Ligation reactions are inefficient when using an insert which has been digested by SalI and SmaI. 

The inefficiency results from the difference in the overhangs.  The four-base overhangs left by SalI ligate approximately 50 times faster than the blunt ends left by SmaI.  The predominant ligation products are:

1. dimers of inserts which have ligated only at their SalI ends. 

2.  dimers of vector which have ligated at SalI ends.  

Ligation of a single insert into a single vector is much more efficient if the two DNA ends have similar lengths of overhangs.  So, if your experimental constraints allow you to use a pair restriction enzymes that leave the same length of overhang, then that is the preferred method.  

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Topic #2:  Improving efficiency of digestion of the pIRES vector:

If your experimental constraints require using an insert which has  been digested by SalI and SmaI then there are methods to improve efficiency.  

On the pIRES vector, the SalI site lies between two SmaI sites in the MCS B region.  But the SalI site is asymmetrically located. It is much closer to one of the SmaI sites.  This asymmetry can be used to your advantage.  

First, digest the vector to completion with SalI.  The resultant linearized vector will have two SmaI sites:

1.    One of the SmaI sites will be extremely close to the end of the DNA fragment.  On one strand, this SmaI site will be separated from the DNA end by a 4-base single-stranded overhang.  On the other strand, this SmaI site has no extra bases separating it from the DNA end.  That closeness to the DNA end will slow the rate of digestion by SmaI.

2.  The other SmaI site will be separated from the other DNA end by 11 base pairs.  Cleavage by SmaI at this site will be  more efficient than at the other SmaI site.  

So, if you perform a minimal digestion of the pre-linearized vector with SmaI, then the predominant reaction product will be digestion at the SmaI site which is further from the DNA end.   

But the SmaI digestion should be performed using a small amount of enzyme for a brief time (i.e., less one unit of SmaI per microgram of DNA for one hour.)  If too much SmaI is added to the reaction (e.g., 20 units per microgram of DNA for one hour) then complete digestion may occur at both SmaI sites.  

After digestion of the vector by both SalI  and SmaI, dephosphorylation of the vector  ends is recommended.  This dephosphorylation is especially important to prevent recircularization of vector on which no SmaI digestion has occurred. 

-paul

If you have the pIRES MCS B Vector the easiest would be to change strategy. If you want to do the partial digest, go for SalI first, afterwards the partial digest with SmaI for different times as mentioned above. Attention: SmaI digests best @25°C, when you use 30°C instead of 37°C, your activity is higher (SmaI is inactivated at 37°C within 15min.) You will be anyway in favour of the SmaI site which is not so close to SalI. If this is the orientation you want you are most likely good. Still will leave you with the problem of coherent-blunt ligation. Dephosphorylation would largely improve the chance of success. FastAP from Thermo is quite nice as it is fast and you simply can heat inactivate before ligation. Another possibility is to use XmaI instead of SmaI (isoschizomer producing 5' overhang.) 

Nikhil, with 25 year's cloning experience, I can guarantee you that the easiest way to accomplish your task is to do a "triple ligation." Cut the vector with EcoRI and SmaI and gel purify the backbone fragment. Cut with  EcoRI and SalI and purify the other backbone fragment. Digest your PCR fragment with SmaI and SalI. Add all three fragments to a single ligation reaction. Transform E. coli. The only way you will get colonies is if all three fragments ligate. Efficiency is greatly improved if you use electrtransformation. I Have made many, many plasmids using this method and have even had success with "quadruple ligation."