I think these are the most likely issues.

1. The most common point of failure I see for cloning gel extracted products is the gel purification itself.

If the DNA gel is illuminated with UV, it must be done at low power, longer wavelength, and the band cutting should be done very quickly. Some UV transilluminators have two wavelength settings, one shorter and one longer. Some have a lower intensity setting for gel extraction.

It's possible for the plasmid to accumulate so much UV damage while cutting the bands that it cannot replicate in E. coli and you will very few or no colonies. If you have access to a dye that can be illuminated with blue light instead of UV, like SYBR Safe, and have access to a blue light transilluminator, this is a better option for gel extraction. The blue light does not damage the DNA to any notable extent and you can take your time working on cutting out bands.

If you are using a column kit to extract DNA from the gel, add 2-3 extra washes of the de-salting buffer (the last wash before elution) and allow the de-salting buffer washes to sit on the column for 5+ minutes each time. In my experience, most kits under report the actual amount de-salting it takes for high quality DNA, probably to seem like their protocol is quicker than competitors. These kits use high concentrations of chaotropic salts to dissolve the agarose, and it's easy for those salts to remain stuck to the silica membrane and carry over into the elution. This can result in very low 260/230 ratios by UV spectrophotometry (ie. nanodrop) which results in inaccurate DNA quantification after elution AND the chaotropic salts will inhibit ligation reactions. This can also be mitigated to a certain extent by cutting out smaller gel chunks and removing extraneous agarose, thus lowering the amount of dissolving buffer that needs to be added to dissolve the gel. This is helped by using blue light as discussed above, because you can really take your time with it.

2. The competent cells may have poor transformation efficiency.

After a failed transformation, it is a good idea to transform an aliquot of competent cells with good quality purified plasmid DNA that has not been treated by restriction enzymes. In this case use your undigested miniTurbo_NLS plasmid. Use a specific amount of plasmid DNA for transformation reaction, usually amounts of 10 ng, 100 ng, or 1 µg of plasmid are used for ease of calculation later on. Choose an amount that will leave you plenty of left over plasmid DNA to work with, though. Do serial dilutions from your recovery (typically no dilution, 1:10, 1:100 and 1:1000 dilutions are enough). When you plate, put the same amount of each dilution onto every plate (100 µL typically). The next day, count the amount of transformed colonies on a dilution plate where individual colonies are easy to count, but you generally do not want to count off a plate that has

Additionaly to what Alexandra answered, I would before try these steps:

• Make sure BOTH your enzymes really work and the vector has the stated sites. To do this, single-cut the vector with EcoRI and XhoI only in separate tubes in the same buffer you use to observe linearization in both cases.
• Check if your PCR primers contain a few additional bases at 5` before RE sites. EcoRI tends to undercut if the site is just at the end. Consult with: https://www.neb.com/en/tools-and-resources/usage-guidelines/cleavage-close-to-the-end-of-dna-fragments for future design.
• After you do step 1, try excluding the dephosphorylation step. The ends produced by your enzymes are incompatible, hence no ligation between them would be expected (if only both enzymes really cut well). But remaining vector phosphorylation could boost ligation efficiency.
• Although this is rare in my practice, yet some enzymes won't eliminate properly after gel, sticking to DNA in a band, thus recutting your ligation products. Inactivate enzymes in your PCR product digestion with heating.

I hope this helps you