6 replies to this topic

[malli]

Hai everyone,

I am working on cloning of ALS genes from last 3months…. i am giving the details of my expt........   
Isolated genomic DNA from leaf samples and amplified with ALS specific primers (350bp).
Ligation: 2µl DNA + 2µl TA cloning PCR 2.1 vector+ T4 DNA ligase 1µl+10x buffer 1µl+ ddwater 4µl kept at temprature14 overnight.
Heatshock with competent (INVαF) cells at 37 for 45 secs. Recovered the cells with 250µl SOC @37 for an hour.
Plated on LB+ ampicillin 100µg/ml and incubated at 37
Observed nice white and blue colonies…..some of the white colonies from each plate straked on new LB+ ampicillin 100µg/ml plates and incubated at 37.

Kept 3ml culture LB+ 3µl ampicillin 100µg/ml and picked a colony from straked plates and incubated at 37. The growth of the cells was not very nice………..Isolated plamid with eppendorf fast plasmid mini kit……noticed nice concentration……..
Did PCR for the miniprep  plasmids with M13 primers. I got amplification at 250bp rather than 350bp (my insert size).
I digested the plasmid with EcoR1. ddwater 4.8µl + 10x buffer 2.5µl+ BSA 0.2µl+DNA 2µl+EcoR1 0.5µl @37 for 3hrs. I run 1% as well as 2% gels with 10µl restriction digested samples. I didn’t notice two bands. (My vector size is 3.9kb, insert size 350bp). I could notice the band intensity difference between self ligated plasmid and plasmid from my samples (too bright and bigsize than control)…………
I run PCR for the prep plasmids with my original primers; I noticed 350bp only in one sample. How can I confirm that my insert is there in the clones…………Please suggest me how I can trouble shoot my problem…….

[phage434]

It sounds as if you have one clone that is right.  That's all you need.  Try cutting again and loading more DNA on the gel.  The 350 bp band will be 10x dimmer than the 3500 bp plasmid backbone (10x less DNA).   So, you need a lot of the plasmid loaded onto the gel to see that band.  You may be able to more easily see the difference between the 3900 and 3500 bp backbones, if you have a way of cutting your plasmid only once.

[murthy]

I think for topo cloning you dont need to use ligase and buffer. please check the manual

malli, on Jul 20 2009, 09:50 PM, said:

Hai everyone,

I am working on cloning of ALS genes from last 3months…. i am giving the details of my expt........   
Isolated genomic DNA from leaf samples and amplified with ALS specific primers (350bp).
Ligation: 2µl DNA + 2µl TA cloning PCR 2.1 vector+ T4 DNA ligase 1µl+10x buffer 1µl+ ddwater 4µl kept at temprature14 overnight.
Heatshock with competent (INVαF) cells at 37 for 45 secs. Recovered the cells with 250µl SOC @37 for an hour.
Plated on LB+ ampicillin 100µg/ml and incubated at 37
Observed nice white and blue colonies…..some of the white colonies from each plate straked on new LB+ ampicillin 100µg/ml plates and incubated at 37.

Kept 3ml culture LB+ 3µl ampicillin 100µg/ml and picked a colony from straked plates and incubated at 37. The growth of the cells was not very nice………..Isolated plamid with eppendorf fast plasmid mini kit……noticed nice concentration……..
Did PCR for the miniprep  plasmids with M13 primers. I got amplification at 250bp rather than 350bp (my insert size).
I digested the plasmid with EcoR1. ddwater 4.8µl + 10x buffer 2.5µl+ BSA 0.2µl+DNA 2µl+EcoR1 0.5µl @37 for 3hrs. I run 1% as well as 2% gels with 10µl restriction digested samples. I didn’t notice two bands. (My vector size is 3.9kb, insert size 350bp). I could notice the band intensity difference between self ligated plasmid and plasmid from my samples (too bright and bigsize than control)…………
I run PCR for the prep plasmids with my original primers; I noticed 350bp only in one sample. How can I confirm that my insert is there in the clones…………Please suggest me how I can trouble shoot my problem…….

[Warren]

Invitrogen has a TA version (non-TOPO) of pCR2.1 as well....

A couple of things.....you made a 10 ul restriction digestion, yet used 2.5 ul of 10x buffer, you should use 1 ul.  For small inserts I usually use 5 ul of miniprep, 1 ul 10xbuffer, then whatever enzyme(s) and water to 10 ul and load the whole thing in a single well.  Usually this will be enough to see the small insert.  Visualizing inserts of that size is additionally complicated by the fact that one of the dyes usually used in loading buffers migrates about that same size and makes seeing it even harder.  If you have a gel-doc system you can overexpose the image and try to visualize bands in that area.  Warren..

murthy, on Sep 8 2009, 10:48 AM, said:

I think for topo cloning you dont need to use ligase and buffer. please check the manual

malli, on Jul 20 2009, 09:50 PM, said:

Hai everyone,

I am working on cloning of ALS genes from last 3months…. i am giving the details of my expt........   
Isolated genomic DNA from leaf samples and amplified with ALS specific primers (350bp).
Ligation: 2µl DNA + 2µl TA cloning PCR 2.1 vector+ T4 DNA ligase 1µl+10x buffer 1µl+ ddwater 4µl kept at temprature14 overnight.
Heatshock with competent (INVαF) cells at 37 for 45 secs. Recovered the cells with 250µl SOC @37 for an hour.
Plated on LB+ ampicillin 100µg/ml and incubated at 37
Observed nice white and blue colonies…..some of the white colonies from each plate straked on new LB+ ampicillin 100µg/ml plates and incubated at 37.

Kept 3ml culture LB+ 3µl ampicillin 100µg/ml and picked a colony from straked plates and incubated at 37. The growth of the cells was not very nice………..Isolated plamid with eppendorf fast plasmid mini kit……noticed nice concentration……..
Did PCR for the miniprep  plasmids with M13 primers. I got amplification at 250bp rather than 350bp (my insert size).
I digested the plasmid with EcoR1. ddwater 4.8µl + 10x buffer 2.5µl+ BSA 0.2µl+DNA 2µl+EcoR1 0.5µl @37 for 3hrs. I run 1% as well as 2% gels with 10µl restriction digested samples. I didn’t notice two bands. (My vector size is 3.9kb, insert size 350bp). I could notice the band intensity difference between self ligated plasmid and plasmid from my samples (too bright and bigsize than control)…………
I run PCR for the prep plasmids with my original primers; I noticed 350bp only in one sample. How can I confirm that my insert is there in the clones…………Please suggest me how I can trouble shoot my problem…….

[HomeBrew]

Warren, on Sep 8 2009, 01:28 PM, said:

Visualizing inserts of that size is additionally complicated by the fact that one of the dyes usually used in loading buffers migrates about that same size and makes seeing it even harder.  If you have a gel-doc system you can overexpose the image and try to visualize bands in that area.

If this is a problem, you can switch to Orange G (see the thread here).  We did -- for just this reason -- and use Orange G loading dye exclusively now...

[Warren]

HomeBrew, on Sep 8 2009, 03:41 PM, said:

Warren, on Sep 8 2009, 01:28 PM, said:

Visualizing inserts of that size is additionally complicated by the fact that one of the dyes usually used in loading buffers migrates about that same size and makes seeing it even harder.  If you have a gel-doc system you can overexpose the image and try to visualize bands in that area.

If this is a problem, you can switch to Orange G (see the thread here).  We did -- for just this reason -- and use Orange G loading dye exclusively now...

hey thanks for the tip, what does orange G migrate as, ie what fragment size does it approximate?  Warren..

[HomeBrew]

Orange G runs at about 50 bp.