Dear Xin Pin,

The selection of your competent cells often depends on their transformation efficiency and yield of the plasmid for your downstream procedures. Is your plasmid a high copy or low copy plasmid type? What is the method of your transformation? Heat shock? Electro proration? Or other method? What is the conditions for your transformation? You need to know that your method of transformation and its conditions may affect the transformation efficiency?

Regard to your ligation, it is good to use high molar insert to vector ratio to minimise the probability of possible vector self ligation. Are you performing in vitro ligation? For enhancing the efficiency of transformation, I suggest you to clean up your ligation products prior to transformation. You can use either column-based techniques or ethanol precipitation for purification of your ligation product. You can quantify the DNA from purified ligation products using Nanodrop.

Depending on the type of your plasmid, high copy vs low copy, you can transform 10-50 ng of plasmid into 50 uL of competent cells.

High copy number plasmid, Heat shock method and what is in vitro ligation and in vivo ligation?  The ligate PCR product is purified or can I take the amplified PCR product directly from the tube with out band excision?

What is the recommended plasmid DNA concentration for transformation say I have two cases where I have least 10ng and a concentration of 300ng of purified plasmid DNA what is the ration of competent cells and ligated plasmid DNA

About ligations, when cloning PCR products a molar excess of the insert over the vector is commonly used. I normally use a molar  ratio around 8/1. 8 molecules of insert per each vector molecule. Others use 5/1 or 10/1. Depending on your insert and vector sizes and of their concentrations in your preps, you can calculate the required volumes. For difficult cloning or if very high ligation efficiencies are required,  its good to try several ratios to determine the optimal one. I know some people does not quantify and just add a fixed volume of vector and insert, but then its a totally blind procedure.

About transformations, the ratio between DNA and competent cells used by different researchers varies in a wide range. If you are transforming with a purified plasmid, this is not very critical. I normally use 50 microliters of cells and 1 microliter of DNA, without taking into account the concentration. For transformation with ligation products, I do the ligation with 100 ng of digested plasmid and the required amount of  insert in a final volume of 20 microliters, then I transform 50 microliters of cells with 10 microliters of the above reaction.

After you do several ligations/transformations, you will find conditions that work in your hands (in your lab with the competent cells you have) and you will use them routinely, These conditions can be different from the ones used by other researchers at other places, but it doesnt matter.

When excess molar of an insert used it could lead to concatamers? although in theory, it wouldn't happen. Also, 1:1 of an insert to vector only works, in theory, I do agree insert ratio should be slightly higher than vector.

yes, we never quantify insert concentration we add 10ul PCR product to 100ng vector and then when I come across insert to vector ratio to be min 5:1 I'm unable to estimate what ration I followed earlier.

With purified plasmid, 50ul cells + 1ul plasmid (51:1) what if I'm asked to estimate the concentration of the cells used in the above reaction? how do I know it? based on OD value?

For transformation with ligation products: if you are taking 100ng meaning 2ul (50ug/ul) and 18ul insert (purified PCR product). Final volume 20ul. Have you calculated the required amount of insert or it's roughly taken?    

and for the transformation 50ul cells + 10ul ligated product (6:1) irrespective of concentration?

Finally, how do I estimate competent cells concentration? Is there any specific paper or book you would recommend.

Using an excess of the insert is quite common and successful, despite the theoretical risk of forming concatamers in some cases and of inhibit ligation by blocking both ends of the digested vector with two insert molecules. How big is the required excess varies from one ligation to another. For instance, when purifying the insert from a gel after digesting a donor plasmid with two non-compatible sticky end-forming restriction enzymes, you can be sure that the insert is perfectly digested and I use 1/1 or 2/1 ratio. For PCR inserts I use 8/1. But if you want to find the ideal ratio in your case, you need to test several conditions.

You can estimate cell concentration measuring the OD at 600 nm. You can find in the web the correspondence of OD and cell concentration for your bacterial strain. Actually I did it only the first time I made competent cells many years ago. Most people never measure cell concentration. I just take 50 microliters of the competent cells to guarantee, although I know this is an excess and lower volumes are also useful. As long as it works, you dont need much more information.

I never ligate by taking  fixed volumes of the vector and insert. So I know the exact amounts I use. I take 100ng of the vector (the volume depends on the concentration of my prep) and the calculated ng of insert  to reach  the desired ratio, for instance 8/1 (again the volume depends on the concentration of my prep). I also add T4 ligase and concentrated T4 ligase buffer and the required water volume to complete 20ul. I take 10 ul of the ligation mixture to transform 50 ul of competent cells.

"For instance, when purifying the insert from a gel after digesting a donor plasmid with two non-compatible sticky end-forming restriction enzymes, you can be sure that the insert is perfectly digested and I use 1/1 or 2/1 ratio. For PCR inserts I use 8/1."

applying the same logic if my amplified PCR template has two different restriction sites on 5' ends then 1:1 or 1:2 works? and PCR inserts with out RE sites only adenine overhang you go with 8:1? why such a huge excess of insert?

"I take 100ng of the vector (the volume depends on the concentration of my prep)" our vector is commercially purchased if at all it is purified by prep I may need to find the cocncetration of the vector.  

"the calculated ng of the insert to reach the desired ratio" so you purify the PCR product from gel or could we take the amplified PCR product directly? from the tube and estimate the double standard DNA concentration (with out dilution/with dilution) and add required volume of insert.  

For PCR inserts you dont have 100% digestion. Digestions are never perfect and digested PCR products usually cannot be separated from their non-digested counterparts (not even using gel purification) due to a very similar size, so your insert is a mixture of double digested, simple digested and non-digested material...When you cut the insert from a donor plasmid the insert band HAS to be digested at both ends. Non-digested products do NOT migrate together with the insert band.  So you cannot apply the same logic to both cases, they are totally  different.This is the theoretical explanation, but I routinely use 8/1 after testing different ratios in several ligation experiments and finding that between 5/1 and 10/1 most of them are ok. You can determine your own ratio, of course. When I need a very high effciency (for library construction), I try several ratios at small scale before choosing one for the whole library. h For PCR inserts w/o  restriction sites like yours, I would definitely try several ratios, because I have less experience with them. This is not difficult, just try a few ratios between 1/1 and 10/1.

You can measure the concentration of any DNA sample, commercial or not, with a nanodrop or estimate it from a gel. 

I always clean-up PCR products using Qiagen columns. You can purify them from a gel, but I never do that, unless the PCR renders more than one strong band.If there is a single strong amplification band (of the expected size) clean-up is enough for my applications.

"For PCR inserts you dont have 100% digestion" I'm talking about inserts that are amplified with restriction sites at 5' ends of primers. So the PCR product has overhangs after it has been amplified. It should be having two different cohesive ends at the amplified PCR product ends that can be readily ligated into a linearized vector.

"donor plasmid with two non-compatible sticky end-forming restriction enzymes, you can be sure that the insert is perfectly digested and I use 1/1 or 2/1 ratio".

Applying same logic in this case, can I use 1:1 or 1:2 vector: insert ratio is what I was asking.

And when you say 'donor plasmid' what exactly does that mean?  

"Digestions are never perfect and digested PCR products usually cannot be separated from their non-digested counterparts (not even using gel purification) due to a very similar size."

Maybe you are talking about digesting a plasmid that has been ligated with an insert? right on the gel, there wouldn't be such high resolution to distinguish them but I'm sure there will be plenty of perfectly digested insert as well so you could purify the insert by extracting it from the gel.

PCR 'clean up' and 'purify' are they both different?  "I always clean-up PCR products using Qiagen columns" you are referring to gel extraction or purifying it directly after amplification?

With ref to this "For instance, when purifying the insert from a gel after digesting a donor plasmid with two non-compatible sticky end-forming restriction enzymes, you can be sure that the insert is perfectly digested and I use 1/1 or 2/1 ratio. For PCR inserts I use 8/1"  

its contradicting "you can be sure that the insert is perfectly digested" but then you say "Digestions are never perfect"? and can't be resolved on the gel.

Case 1. Yo amplify the insert by PCR with primers containing restriction sites and overhangs. Digestions are never perfect and digested and non-digested insert have similar migration patterns, so they cannot be separated. Use a big insert excess for ligation. The optimal excess can be determined trying several ratios.

Case 2. The insert is cloned in vector 1 (donor) flanked by two restriction sites. You digest the insert from this vector to clone it in a second vector using the same restriction sites. Digestions are never perfect, but non-digested products migrate much less than the digested insert (if the insert is quite smaller than the donor vector). So the only product migrating at the insert size is the perfectly digested insert having both sticky ends. After gel purifying, you don't need such an excess of the insert for ligations.   

Again, case 1 and 2 are different, It makes no sense to apply the same logics. There is no contradiction. My answer is exactly the same.

I always clean-up PCR products using Qiagen columns without gel extraction, unless there is more than one strong amplification band. In the latter case, its necessary to use a gel to isolate the desired band.