Depends on concentration, if you have detectable amount of contaminating DNA in your Sybr, than yes. But you should be able to find that out by exciting a stock of sybr you are thinking is contaminated and fresh not opened before sybr and check the fluorescence.

Maybe you can try berberine or hydroxynaphthol blue stains, you can add them before reaction:

 http://www.biotechniques.com/BiotechniquesJournal/2015/April/Shining-a-light-on-LAMP-assays--A-comparison-of-LAMP-visualization-methods-including-the-novel-use-of-berberine/biotechniques-357796.html

HNB works well in my lab, berberine we are going to test.

That's a great suggestion, I will try that. I've used HNB in the past but could never get it to work. Are using a concentration of 120um in your final 25ul reaction?  I never saw a colour change between positive and negative samples and my starting colour was never purple but more of a dark blue, We tried titrating it alongside Mg2+, to no avail as well. I haven't tried berberine yet but I may if I can't get the other dyes to work. Thanks for your help!

Yes, we use HNB at 120 uM final conc. This is strange, in my lab this works well for different viral pathogens. I am attaching result for dilutions of sap from plant infected with Potato virus Y. The sky blue are positive and dark blue are negative samples (sap from healthy plant and last - right tube is mock - water added instead of sap).

Maybe you should try to use pre-made kits containing fluorescent stain to check if reaction is going properly. We routinely use kits from OptiGene sold by Novazym Poalnd:

http://www.novazym.pl/isothermal-master-mix-optigene-c-276_267_127_188_149.html

We just add primers and template and run on real-time machine using FAM chanel to monitor fluorescence increase during amplification. This works very well and is our control for colorimetric assays we try to develop.

There are two possibility that negative control tube shows positive. First is contamination as you are concerning, and second is nonspecific amplification by primer dimerization. LAMP assay is well known as specific DNA amplification method, but LAMP primer set occasionally cause nonspecific amplification. Therefore I recommend you to analyze negative control tube which shows positive by gel electrophoresis. If it was same band pattern with positive reaction, it is contamination. If not, including smear pattern, it is nonspecific reaction like primer dimerization.

Totally agree with the Yuki's answer.  I strongly consider the contamination in LAMP is primarily caused by the primer-derived template-independent non-specific amplification. Actually LAMP is a mixed amplification mechanism which contains the excepted LAMP scheme and the template-dependent linear target multimerization and amplification (LIMA). I think LIMA is caused by low-fidelity Bst DNA polymerase. The other form of LIMA is template-independent, and its efficiency greatly relies on the stability of primer-derived duplex structure that is also the commonly called primer dimer. Therefore, to address the issue, two measures may be advisable. One is to check your primers on whether there's a strong duplex structure formed. If so, redesign your primers. Alternatively, I recommend a simply way to do it. Focusing on the negative controls, pick out the two, three, four, or five primers from the six LAMP primers to do PCR-like LAMP, and then you'll find which primer or  primer composition that can bring about the non-specific amplification. Subsequently, only modify and redesign the "troublemaker" until the problem is solved. The other is to set a proper reaction time, since it is sure that a long reaction time greatly challenges the Bst polymerase's loyalty. How to define the proper time is also easy. Do a plenty of negative controls in intra-batch and inter-batch assays and a time shorter than the time of negative controls is the one you need. Usually, 60 min is mostly proper for LAMP.  Hope it useful and always free to contact me with further communication.