1.6 kb might be too long for Taq pol, try to extend the extension time, I would suggest 3 min. Alternatively, try a different DNA pol such as Phusion DNA pol.

Plasmids are coiled and supercoiled so can be hard to amplify. You could try restriction cutting the plasmid with an enzyme that does not cut your insert. You appear to have a lot of primer dimer which will make th pcr reaction fail so I would use much less primer and ,ideally, a hot start enzyme (polymerase). Also I would use much less plasmid dna--it is a small molecule and use a 20ul reaction volume as larger volumes are more forgiving to small changes in reagent concentrations and the evaporation of some of a small volume means that some reagents will be too concentrated. Measure the od of your primers.It is eadier to have an accurate amount of primer if you know the concentration accurately and it is likely that you are using far too much primer

Thankyou Libor Krásný and Paul Rutland for your valuable insights.

I will try doing the PCR in a 20 ul reaction as well.

I actually did a serial dilution of the primers and figured out what dilutions to take based on band intensity of the primers.

This helped me reduce the problem of primer dimers. I also switched to Phusion since I anyway need high fidelity amplification.

When I did my trial PCR in Phusion Pol, I found that I am getting an additional band (of 1 kb) apart from my amplicon which is 1651 bp.

I have two images - one short and one longer run.

Extension time of 30 s

Annealing time of 35 s

Number of cycles : 35

DMSO 3% final in 10 ul volume

left to right:

lane 1: lambda ladder

lane 2: no template control

lane 3: 10 ng plasmid, Ta = 49 C

lane 4: 16 ng plasmid, Ta = 53 C

lane 5: 6 ng plasmid, Ta = 53 C

lane 6: 16 ng plasmid, Ta = 53 C

lane 4: 10 ng plasmid, Ta = 55 C

These are all 10 ul reactions set as per NEB Phusion Polymerase protocols.

Instead of 1:250 for Forward and 1:25 for Reverse, try serial dilutions to find the optimal reverse primer concentration.

Use equal molar concentrations (e.g., 0.2–0.5 µM each).

Prepare new primer dilutions with accurate pipetting to ensure reproducibility

Run a gradient PCR from 52°C to 58°C to determine the best annealing temperature.

11 ng in a 10 µL reaction is a bit high for plasmid DNA. Try reducing the template to 1–5 ng to prevent excessive background signal.

You have a total of 4.0 mM MgCl₂ (1.5 mM in buffer + 2.5 mM added), which may be excessive. High Mg²⁺ can promote non-specific amplification.Try 2.5 mM total first (reduce supplemental MgCl₂ to 1.0 mM).

Taq polymerase extends at ~1 kb/min, so 1 min 40 sec is borderline.Increase extension time to 2 min to ensure complete amplification.

Reduce primer concentration if needed (e.g., start with 0.2 µM).If primer-dimers persist, try adding 5% DMSO or betaine to improve specificity.

6-16 ng is WAY too much plasmid DNA for PCR. You only need 0.1-1 ng.

As my advisor said, "you can't rush quality".

My guess is that you are getting 0 amplification and just seeing the plasmid on the gel. 10 nanograms is just enough to be visible on a gel when you use ethidium bromide. And wash your gel combs - that ladder is really smeared.

Step 1: Find a restriction enzyme that will make cuts in the plasmid, but not in the region you want to amplify. If possible, get one that's heat-inactivated so you don't need to do a cleanup.

Step 2: Do the math so that you can add in a reasonable amount of digested plasmid DNA in your PCR. You want to use 0.1-1 ng in the PCR so dilute accordingly. And you'll want the volume of diluted plasmid to be one you can reliably measure (2-5 microliters is a good range).

Step 3: order primers and use a recommend concentration.

Step 4: use your diluted, digested plasmid with the Phusion polymerase to set up PCR. Use 20-50 microliter reaction volumes. 10 is too small a volume to measure each component accurately.

Good luck!