Hi there, 

Have you performed any purification after your PCR? like some PCR clean up kit. If you did not purify your sample, I think it is possible that your sugar might have trapped your DNA which delay its migration rate as the sugar does not carry charge. Hope that helped you

I agree with Chun's assessment as contamination being a possible source of the problem. I would also consider modifying (lowering) the concentration of your agarose gels to see if that has any effect, as well as voltage and run time. Just load a couple samples to evaluate the changes seen from altering the protocol, and if results are improved then repeat with all your samples.

Below is a description of the method/protocol I used back when I had to run a bunch of gels. I was assessing DNA purity and fragment size, post-sonication and pre-PCR (target size was ~100-150bp, the accuracy/validity was crucial for me bc methylation capture performed downstream would be compromised if fragment size/purity was not assessed correctly on GE) 

Hope this helps!! 

Protocol:

Agarose gels were mixed to a concentration of 1.75% with the following components: 0.445 grams of agarose powder dissolved in 32 ml dH20, 8 ml 10× TBE (1× final), and 0.8 μl ethidium bromide (EtBr). The liquid agarose was poured to a depth of 6 mm in a clean 5 × 8 cm casting tray and allowed to cool for 30 minutes. Running buffer (1× 1M TBE) was prepared with 204 ml dH20, 51 ml of 10× 1M TBE, and 2 μl EtBr.

Upon filling the electrophoresis tank with running buffer an additional 4 μl of EtBr was mixed at the anode terminal. Ethidium Bromide migrates in an opposing direction to that of DNA; therefore this step was performed to increase the potential number of collisions between EtBr and DNA, which effectively increasing the fluorescence of samples.

Stock 10× TBE, used for gel and buffer preparation, was made by dissolving the following ingredients in 1 liter of deionized water: 108 grams Tris-base, 55 grams boric acid, and 7.5 grams EDTA. Lanes were loaded with a 10 μl mix that consisted of 500 ng DNA, 2 μl of 5× sample buffer (Bio-Rad Laboratories, USA), and nuclease-free H2O used to adjust the final volume. Samples were run for 01:10 hours at constant voltage (100V) and immediately visualized on a UV transilluminator (AlphaEaseFC software; Alpha Innotech, FL, USA). A 100-1000bp molecular ladder was loaded adjacent to samples to assess the extent of DNA fragmentation

looking at the image of your gel a second time, i'm also thinking concentration of your loaded sample could be a factor. I assume you measured DNA concentration prior to mixing samples to load, and normalized the amount of DNA loaded by adjusting the volume of each sample... 

I would expect to see those results on your gel if the concentration was off the charts high, you had a contaminant, or there is an conflict between sample/gel interaction...

Hi, I'm just wondering whether or not there is a small amount of contaminating DNA in your maltodextrin sample that when spiked with your control DNA unfortunately causes concatemerisation during PCR.  In this case your qPCR probe would still be cleaved during amplification but would not result in discreet 100 bp fragments, rather a range of larger size fragments as seen.  This however doesn't explain why you're not getting a 100 bp band in your DNA + Maltodextrin pre sample.  What is the essential difference between this and the maltodextrin spiked pre sample? Should they not be exactly the same thing +/-  the PCR components? I presume you're using a hotstart taq in whichever qPCR kit you're using?

It looks like that the 'procession (of DNA and maltodextrin)' caused this problem.

1. We don't know how did you 'process' them. The 'procession' seemed to 'link' the DNA  to the maltodextrin (polysaccharide). And maltodextrin could be too large to swiftly move around on the 4% gel. So the image showing the stuck DNA ('linked' to the maltodextrin) around the well.

2. On the other hand, the gel showed that your spiked sample (maltodextrin + spiked DNA) had no such problem, because even they co-exist, but not 'linked'. So when you electrophoresed them, the DNA could still freely move down the gel while the maltodextrin stuck in the well.

3. Besides, it looks like that all the 'spiked DNA' (sample of maltodextrin +spiked) were freely moved down, because the lane of 'water+ spike DNA' has same DNA band intensity as the lane of 'maltodextrin+ spiked with DNA sample).

4. Seems that maltodextrin inhibited PCR reaction, because the band intensities on gel for '[(maltodextrin + spiked PCR) (Pre)] and [(maltodextrin + spiked DNA) (Post)] was similar (if the same volume of DNA solution for both was used to run the gel). However, it seems that PCR inhibition did not occur on the 'processed (maltodextrin + DNA)' samples, because the band intensity of (Post) is much brighter and larger than that of (Pre).

5. How come your 'maltodextrin only (post)' has two faint PCR bands? Primer dimer?

You can adjust the concentration of the agarose to give bigger pores so bigger sample can migrate. This can be a possible reason while your sample didn't migrate in 4% agarose. If you decrease the concentration the sieving of the gel will increase allowing bigger molecules to migrate. However be careful because small molecules can then migrate much quicker so you would need to run the gel for a shorter amount of time.