Hi everyone,

Would appreciate any help with this problem.

I have established in-lab protocols that have seemed to work in extracted and detecting eDNA from water systems (we have fish systems in lab, both freshwater and saltwater). These methods have also worked in semi-representative environments of the ocean conditions such as detecting eDNA in holding tank water from facilities that hold the marine invertebrate we are trying to detect (they also use ocean water directly off the coast in their systems). However whenever we try to apply these same methods to field samples (surface water samples near a kelp forest), we cannot detect any environmental DNA. Even when using general primers such as 16S, or more general universal barcoding genes that are expected to detect at least something. The nanodrop readings for these DNA samples also look fine, with a A260/A280 ratio that usually ranges from 1.8-2 and usually above 20ng/ul. My thought is that inhibitors are present in these samples, as even when I spike these samples with clean and high yield DNA (in PCR) they still do not show detection.

If anyone has encountered seawater PCR inhibitors before, or maybe believes that it could be something else going on please let me know and I appreciate the help!

General Methodology Below.

Methods we are using,

1.) Filtration of water (1-3L, any volume in that range seems to work so far) through a .45uM nitrocellulose filter.

2.) Extraction of eDNA from that filter using a DNAeasy power water kit (following the protocol exactly).

3.) Then amplification of this DNA using PCR.