This is about detecting the activity of a splice-modifying oligo by reverse-transcriptase PCR and gel electrophoresis.
The expected outcome of an exon targeting an exon 3 splice junction (either i2e3 or e3i3) is elimination of exon 3 from the mature mRNA (along with both of its flanking introns, i2 and i3). The typical way to assess activity of a splice-modifying oligo is to isolate RNA from treated samples and run reverse-transcriptase PCR (RT-PCR), then run the PCR product on a gel to assess the mass of the product against a DNA ladder.
I have some notes on the expected outcomes of splice-modifying oligos (and a description of some possible unexpected outcomes) elsewhere on my blog:
To detect elimination of exon 3, we would typically design primers to exons 2 and 4, set back from the e2i2 and i3e4 junctions sufficiently that the RT-PCR product without exon 3 will still be about 100 bases long (that way enough dye binds in the RT-PCR product to see it clearly on a gel; that's tough if the fragment is too short). For short exons, it might be necessary to place the primers more distantly, for instance in exons 2 and 5. If the product is subject to nonsense-mediated decay, you might not see the splice-modified band; instead, NMD is visualized as a dimming or disappearance of the band when compared to the wild-spliced (negative control) band. To compare band intensities, load the wells lightly so the bands are not saturated and compare with RT-PCR of a housekeeping gene to confirm your total RNA loading is comparable between wells. If the wells are overloaded (so the bands are saturated), you might not see a partial knockdown of the wild-spliced band.