Based on your data, there is no way to know if your primers are useful.

You need to measure the efficiency of your primers before you do ANY more experiments. See this website for a good introduction.

https://www.thermofisher.com/us/en/home/life-science/pcr/real-time-pcr/qpcr-education/absolute-vs-relative-quantification-for-qpcr.html

Adding to Katie's comment, I'd suggest digesting your RNA prep with DNase before reverse transcription. Residual genomic DNA in your RNA prep can be problematic especially when you deal with non- or poorly spliced transcripts such as lncRNAs, because there is no way of designing primers which allow to discriminate gDNA and cDNA, e.g. exonic primers flanking long intron(s)

Hi Wittawin, of course I agree with the answers above. The gDNA problem is of course an issue in your case, GAPDH is useful for this purpose since it has a few pseudogenes in the genome and if you don't remove gDNA efficiently you'll get an amplification. Although a difference of 7 cycles is acceptable (it means your gDNA is around 1% of the cDNA, the exact amount depending on the efficiency of your primers and their linearity range... that's what Katie's suggestion would allow you to know), a Ct of 27 means a quite high amount of gDNA in your sample, that's a Ct typical of a low expressed RNA (mRNA or lncRNA...!), which means it will not allow you to detect it properly. Assuming that the melting curves refer to the same amplicon (and it should be verified, the 3 degree C difference might also mean that in the absence of the specific template you amplify something non-specifically on the gDNA of a very similar length and/or base composition), in fact you have a very small Ct difference for your lncRNA when you omit the RT step, a 2 cycles difference means you are around 25% gDNA contamination, which will heavily affect your relative quantification.

So, to summarize, these would be the suggestions:

1. Obtain a better RNA extraction so that after DNAse treatment your GAPDH Cts are in the range of 32-35 cycles

2. Make a standard curve for the two genes with your primers (serial dilutions, at least 5, of your cDNA: for GAPDH you can dilute 10-fold every time - so to have 1:10, 1:100, 1:1000, etc of your cDNA -, for your lncRNA which might be low expressed better go by 2-fold dilutions - 1:2, 1:4, 1:8, etc... It's an example or a suggetion, since I don't know how much you diluted your cDNA in the experiment you describe. Possibly you can do 5-fold dilutions, which would be better...

3. Verify the efficiency of the amplification with your primers interpolating your results: the standard curve should have a r2>0.990. You might realize that at the higher or lower concentrations (or both if your standard curve is long enough) you lose linearity: that's a very important point, if you don't have linearity, forget the quantification. You have to be in the linear range of amplification to be able to quantify, for both the calibration/reference gene and the GOI... so if the lncRNA is low expressed, GAPDH might not be the best choice to calibrate (too high expression), choose a transcription factor, typically much lower expressed (for example the TATA box binding protein, TBP). And if your lncRNA cannot produce a standard curve (you lose linearity after 2, 3 dilutions), either you have to increase the amount of template (use more RNA to produce more cDNA, use more efficient RT kits or RTs, prepare a cleaner RNA) or try to design more efficient/specific primers...

Good luck!