Hi Hazel,

The best strategy is to have a very good reference so you can design specific primers that are long enough and anneal at high temperatures to avoid your primers going where they're not supposed to be going. I'd recommend designing something not shorter than 25 bps with a melting temperature of 55 C for both oligos. Make sure you get them from a vendor with strict quality controls. IDT in the US makes almost fool-proof oligos whereas Sigma makes quick and dirty versions than fail every now and then. You know how serious QC is when it takes several days to get them ready; if they are in your lab overnight then I wouldn't think they passed all barriers.

A second, more complicated strategy is to have long PCRs that amplify say half of the mitogenome spanning your region of interest, in this case CR. With very long fragments, you can use those new long pieces that should only be of mitochondrial origin as a new PCR template, ensuring the amplicons you get are no longer using the whole genome DNA extraction and thus preventing nuclear copies. This technique of course is time consuming and more expensive. 

Why don't you try making new and very specific primers? That does the trick 95% of the times!

--Hernán

If you are having difficulty distinguishing them from the outset and have access to some soft tissue (liver is ideal), you can isolate mtDNA by differential centrifugation, then redesign primers to be specific to that sequence. As numts have a much lower mutation rate, and there is usually only one amplifying, they are usually easy to identify as a contaminating sequence. They are often useful in themselves as an outgroup for the real mtDNA- the low substitution rate means that you almost have the equivalent of a fossil ancestral genome to work with. So you might want to design primers for the numt itself too.

Hi all

Many thanks for your help so far. I am also working on museum specimens so have designed short fragment overlapping primers. To get these I used some general primers on modern dna samples, then redesigned some species specific primers, then used this sequence data to design my museum primers. 

Does this seem like the correct protocol to avoid Numts in the museum amplifications? 

Thanks again 

What you're doing is the traditional method for getting sequence data from museum specimens. While this in theory is OK, you must keep in mind that molecular work using degraded samples is notoriously prone to contamination. You should try to do most if not all of the wet lab steps -from extraction to PCR- in a different lab with dedicated pipettes, tips, and reagents that haven't touched your modern parrot samples. Otherwise you risk amplifying modern DNA stuck to your regular equipment.

I assume you're extracting from toe pads? Make sure you wash your dissected pieces multiple times over a week period over ethanol cycles at different concentrations, finishing with a full ddH2O overnight wash. Then supplement your digestion buffer with DTT. I'd recommend also using phenol chloroform to extract as much DNA as possible.

Another issue to consider is that amplifying really small DNA fragments (

Hi Hernan

Ok, I should point out here I have already conducted the bulk of the lab work.  I conducted ALL my museum work in a separate museum dedicated DNA lab with very strict protocols to ensure no contamination.  I am also happy to report I used a Hot start Taq.  

Thank you for all your comments so far! Very helpful indeed.