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Environmental DNA - Science topic

Explore the latest questions and answers in Environmental DNA, and find Environmental DNA experts.
Questions related to Environmental DNA
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6 answers
The DNA needs to be pure, clean, free of inhibitors and suitable for quantitative PCR.  The current method we use is limited to 0.25g of soil and I would like to use at least 10g, more if possible.
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I hope this maybe useful:  Plassart P, Terrat S, Thomson B, Griffiths R, Dequiedt S, Lelievre M, Regnier 
T, Nowak V, Bailey M, Lemanceau P, Bispo A, Chabbi A, Maron PA, Mougel C, Ranjard
L. Evaluation of the ISO standard 11063 DNA extraction procedure for assessing
soil microbial abundance and community structure. PLoS One. 2012;7(9)
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1 answer
Hello
Im an MRes student new genetics and qPCR, and am developing a species-specific primer/probe set to detect Najas flexilis, a rare aquatic macrophyte in Scotland. The aim is to detect the plant by eDNA filtered from freshwater. However, I am running into efficiency problems with qPCR. Can you help? If so, firstly, allow me to explain the development steps.
Using Primer3 software, I developed a pair of primers with Tm ~ 53/55 C. The forward primer showed high hairpin stability in silico. After running a gradient PCR, it was discovered that 59 C gave the greatest concentration of amplicon, perhaps because the melt temperature was too high for hairpin formation (?).
The probe that was designed with the primer pair had Tm ~62 C. A qPCR, using Environmental Master Mix (different to the master mix used in the aforementioned gradient standard PCR), was undertaken. The thermal protocol used was based on work for Great Crested Newts, rather than that recommended in the master mix manual, as recommended by my supervisor:
50 C for 5 minutes
95 C for 10 Minutes
Then 55 Cycles of:
95 C for 30 seconds
59 C for 1 Minute.
The qPCR resulted in amplification of eDNA extracted from filtered water, and of a dilution series extracted from a live tissue sample. The efficiency was only 44%. I'm keen to increase the efficiency of the reaction, perhaps by reducing inhibition, or modifying the thermal profile based on the primer/probe set designed. This in turn will improve the potential to detect Najas flexilis using eDNA extracted from freshwater this summer.
Qiagen Soil Kits were used in the extraction of live plant tissue used in the qPCR dilution series, which, like Environmental Master Mix (EMM), helps reduce inhibition. I will soon test for inhibition using an IPC (waiting for it to arrive), but suspect the probe design is the problem, or the thermal cycling condition.
The amplicon is 193 bp, higher than the maximum recommended 150 bp. The high hairpin stability of the forward primer is may still be affecting the amplification.
From what I have detailed above, can anyone recommend next steps to increase efficiency, before I begin analysing samples collected in the field?
I was thinking of running a gradient PCR using Environmental Master Mix and see if the greater concentration occurs at a Tm that is different to 59 C.
I also thought of qPCR dilution series in triplicate, at different temperatures. My instincts suggest that 57 C might be a better Tm, as it is 5 C lower than the Probe Tm.
Also, I thought that perhaps the the thermal cycling could have three steps, as in standard pcr, with a 72 C elongation step added to aid in elongation 193 bp.
In truth, I am unsure, but before delving into the unknown, thought I would ask those who might know.
Any thoughts would be greatly appreciated.
Best Regards
Nicholas Crutchley
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If you haven't done this I would run a primer concentration optimisation by taking 3 concentrations (50nm, 300nm, 900nm) and running PCRs at the 9 potential combinations against a single concentration of DNA. The lowest average CT value would indicate the optimal primer concentration
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3 answers
I would like to construct cosmid library using metagenomic DNA extracted from soil sample. After extraction, I used electroelution technique for eDNA purification. But, I loosed alot of eDNA during this step. In addition, it cannot be ligated with the cosmid vector.
I thought that eDNA might not be pured enought.
Does anyone have suggestions about eDNA purification for metagenomic library construction?
Are there any purification kit available for large DNA fragments?
Thanks in advance,
Apirak Wiseschart.
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Dear Apirak, please check the attachment in pdf format for eDNA isolation and purification protocol. The eDNA preparation method was used for microarray study, however it could be applied for your experiments. please note that the protocol is bit time consuming.
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5 answers
Hello,
I need to develop a cost-efficient method of detecting Ondatra zibethicus and Myocastor coypus presence in various freshwater environments (rivers, lakes,...) so I thought about using eDNA. Since I didn't come across any research papers using the method to detect the above mentioned species, I wonder if it would be efficient?
Kind regards,
Sonja
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That is very helpful, didn't come across it yet, thank you very much!
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4 answers
Hello Everyone,
I am trying to visualize eDNA in S. aureus biofilms but my strains already express gfp constitutively so finding a fluorescent stain that will emit in the blue or red region is very much desired. Any suggestions? Thanks
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Hi
I'm not sure if you'll be able to find a dye with blue fluorescence unless you use UV and it's a bad idea for living biofilms. I'm agree with Alexander, the best options are phenantridiniums.
best regards
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1 answer
Hello everyone,
Recently, I plan to extract environmental DNA/RNA from biofilm. The media is activated carbon. Considering the strong adsorption capacity of activated carbon, is there any good method for the extraction? Any of your help would be appreciated!
Millions of thanks,
Lin
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Dear Sir. Concerning your issue about the method of DNA/RNA extraction from activated carbon biofilm. The biologic activated carbon (BAC) process is widely used in drinking water treatments. A comprehensive molecular analysis of the microbial community structure provides very helpful data to improve the reactor performance. However, the bottleneck of deoxyribonucleic acid (DNA) extraction from BAC attached biofilm has to be solved since the conventional procedure was unsuccessful due to firm biomass attachment and adsorption capacity of the BAC granules. In this study, five pretreatments were compared, and adding skim milk followed by ultrasonic vibration was proven to be the optimal choice. This protocol was further tested using the vertical BAC samples from the full-scale biofilter of Pinghu Water Plant. The results showed the DNAyielded a range of 40 μg·g−1 BAC (dry weight) to over 100 μg·g−1 BAC (dry weight), which were consistent with the biomass distribution. All results suggested that the final protocol could produce qualified genomic DNA as a template from the BAC filter for downstream molecular biology researches. I think the following below links may help you in your analysis:
Thanks
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3 answers
Hi, I'm trying to get environmental DNA from human armpit skin microbiome for metagenome analysis using T-RFLP. Any suggestion for the best product or protocol to get most DNA from these kind of sample? Which DNA swab is recommended?
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You could give Catch-All™ Sample Collection Swabs from Epicentre a try. We have used it for food allergy- skin microbiome study and got decent yield. We used Mo Bio Power Soil kit (now it is Qiagen DNeasy Power Soil kit). Qiagen reps recently recommended QIAamp DNA Microbiome Kit for host depletion and particularly for swabs and body fluids.
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I have done MiSeq sequences on ice samples and I would like to run the analysis for them. Nevertheless the two pipelines I'm familiar with (Mothur and Parallel-META) are not the best for ITS1 analysis. I have look in google to find tutorial (Mothur is absolutely fantastic for this) but I can't find anything similar for Skata.
Help anyone?
Thanks,
Mario
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Hi Mario,
I recommand reading this SOP from LangilleLab:
This should get you started to analyse your ITS data.
Cheers,
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3 answers
Hello,
I'm trying to amplify a gene in both eDNA and bacterial DNA (a strain of Bacillus, that should be the positive control) using degenerative primers, their sequence was published in a (decent) paper.
However, following the exact authors protocol (with the exception of the buffer used)- I'm not getting any bands whatsoever.
I've been trying to do gradient PCR- but still, no bands (67.5 is the authors recommended annealing temperture, I've tried the whole range of 56-72). I found out that the buffer I'm using (Dream Taq) has 20 mM of MgCl, while the buffer that was used in the paper (FailSafe PCR Buffer) has 4 mM of MgCl- could that be the reason?
I currently don't have a positive control, by using the same reagents but dufferent primers, and the same PCR machine yielded efficient amplification, so I believe I'm missing something.
Thansk you.
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Time to start trouble-shooting to find a good positive control. You've learned that your enzyme works, your thermocycler works and you can set up a successful PCR reaction. That's a good start! Try out a different set of primers with your Bacillus DNA to confirm that your DNA extraction worked. Then test out the published primers. I wouldn't think the MgCl matters that much. What are the recommended annealing and extension times?
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4 answers
Please if we want to check many related species in the same eDNA sample we should use more primers or more samples or sequence all genome extracted from samples
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I am so grateful for your reply dear colleagues.
Actually I want to checking the species (belong to same family) in an environment. All of those species barcode by one primer so how we can determine the species composition by using the same primer.
Dear Leah Clarke : Is this technique is efficient to investigate species composition even if they related with the same genus or family. Please reply
Thank you for all of you Dr. Combik, Dr. Clarke and Dr. Laldin.
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Did microbial DNA of sessile bacteria can integrates the genome of tumorale cells ??
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4 answers
I have environmental DNA samples extracted using the MoBio/Qiagen PowerWater kit and quantified using a Quantus fluorometer. While most of my samples have concentrations of >100ng/uL dsDNA, a few have around 15-30ng/uL (in 100uL total volume) which might not be able to satisfy the 2,000ng input DNA required for PCR-free sequencing.
Will it still be possible to submit these samples for PCR-free shotgun metagenomic sequencing (150bp paired-end)? Or would my only option be to re-do sampling with more biomass for extraction?
*I really would not prefer to do PCR-based sequencing or metagenomics because of potential bias introduced by the technique.
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So you are sequencing all extracted material, and not enriching for any marker(s) in particular?
For the samples at 15-30ng/ul in 100 ul, assuming quality is there, I would think these would be fine after concentrating down to a smaller volume.
Could it be an option to use PCR during library preparation, but then collapse PCR-duplicated polymorphisms by identifying them bioinformatically, either using fragment length polymorphism (assuming that you can build contigs- which would then be of variable length), or by incorporating a degenerative region into your adaptor design, thereby mitigating (or at least minimizing) any bias?
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Dear all. I am working on Earthworms and I am trying to recover their DNA from the soil and use the 16S and COI gene to prove that we can use the environmental DNA from the soil to identify different species of epigeic earthworms that are used in the vermicomposting processes. Bienert et al. (2012) forms the basis of my research since they proved that it is possible to track earthworms from the soil using their DNA.
My problem however, is that I am struggling with the PCR protocol to use in order to obtain positive results. Bienert et al. did not provide the protocol in their paper of these two genes and most of the literature has tissue DNA protocol for these genes instead of the eDNA. Please help the complete PCR protocol (reaction mix and PCR setup)
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I think I found a good enough protocol for your purpose. See the attachment here.
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I am doing qPCR (TAQMAN assay) for fish DNA samples. I have extracted DNA by a commercial kit usually using successfully in our lab. Now I want to calculate PCR efficiency for standard curve and it is 2.6 (~144%) with r-square value of 0.99. I don't know why it happened and now what should I do? It would be great if I could get some nice suggestions. thanks
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Struggling with understanding and calculating qPCR amplification efficiency? This article breaks it down for you in an easy and comprehensible way. It also provides a treasure chest of resources that will bring your qPCR game to the next level ->
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Hi, I'm working on metagenomic analysis of undefined mixed culture of algae-bacteria culture using 16s/18s rRNA analysis using Illumina sequencer. Any suggestion for the best kit to do total DNA extraction from algae biomass?
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Hi, We have used MoBio power soil DNA extraction kit and we found few genus belong to Cyanobacteria in our recent studies from sediment samples.. !! However we have never tried it specifically for Algae.
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Or other important methods (outside of standard Molecular Biology techniques) that would be appropriate to have a grasp of if wanting enter the field. 
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Try to play with MG-RAST server and MetAMOS pipeline:
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I usually use distilled water for primer dilution, i never got problem but one of GC rich (63%) primer getting degraded often. which one can be better for GC rich primer dilution TE or water if TE what is the concentration and composition and is there any affect while doing sequencing.
Thank you..
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you should always dissolve pcr primers in TE. Water dissolves CO2 from the air to fprm carbonic acid which results in acid depurination of the primer and degradayion and the alkaline buffering of Tris prevents this. The EDTA chelates divalent ions  like magnesium so this prorects against nuclease degradation of the dna, There will be no effect on sequencing  which is a robust reaction and the small amount of primer added will have no effect on the larger amount of sequencing buffer with magnesium and manganese in it. 10mM TE is plenty to buffer the effect of CO2 absoeption
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I had already quantified my DNA using a Spectrophotometer, and most of the samples were around the 200-300 ng/ul. When I quantify using Picogreen assay, my standards range from 10-1000 ng/ml. I know the conversion is 1000ng/ml= 1ng/ul... So, if one of my samples concentration with the picogreen is 600ng/ml that means I only have .6ng/ul?? 
I have been trying to find out about this, maybe I'm not doing the conversion right? Please, someone help. 
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It is 'very' unlikely that a spectrophotometer can measure 'anything' that low in concentration (not ppb or ug/L or ng/mL).  A ppm (typical for a spec.) is ug/mL or 1000 ng/mL or 1 ng/uL
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4 answers
I need full-proof protocol for isolation of extracellular DNA (eDNA) and its characteristic in gel.
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Dear Subhaswaraj Pattnaik,
I'm not sure if this will help you, but I extracted eDNA from Candida albicans biofilms and the protocol I applied can also be used for bacterial biofilms. However, I didn't evaluate eDNA in agarose gel.
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23 answers
Hi,
I'm trying to extract DNA from P. larvae spores for PCR detection.
I know that there are some attempts (Bakonyi et al. 2003/ Alessandro et al. 2007), but I wonder if there is any comercially available kit to extract DNA from spores.
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Thanks for the reference! I will try with the kit.
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Cataloging frogs calls are a robust method of collecting data for bullfrog immigration, but they don't work year round, are subjective and need dedicated volunteers. eDNA needs specialists and resources. The early methods of airguns for sample collecting are unlikely to be approved for urban public spaces.
Fyke nets have promise, but permission to deploy them is not a given, plus costs, maintenance and perceived impact on ecosystem are problems.
Is there any method that is virtually free, quick and easy? Wondering if there is any experience of identification of species via photographs of individuals like they do for whales?
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Hello,
You can ID individuals based on patterns, and there are softwares that help, but it is quite species dependent. It should work pretty well with the spots on the ventral side for bullfrogs though. The paper below give some recommendations on what works and what is better avoided.
For free methods, I still recommend call monitoring, it works quite well for population dymanics:
Best,
Amael
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1 answer
I am attempting to quantify fungal presence in soil eDNA samples. Using normal PCR (BioBasic master mix) the primer set amplified well, and we optimized for annealing temperature and primer concentration. When we moved to qPCR (PowerUp SYBR green master mix), there was absolutely no amplification. Other primer sets on the same DNA sample amplified well, and even negative controls amplified more than the fungal primers did (very late, at 35-40 cycles). We have repeated this several times, with slightly varying temperatures and cycling parameters, but there is still no amplification.
We have no idea what would completely prevent amplification in qPCR, but not PCR. It doesn't seem to be an issue of efficiency, as at least some amplification would have been seen. The primer set has also been used previously for qPCR, and we have attempted to mimic their procedure. We think it is most likely to do with the contents of the master mixes used, but can't think of what. The main difference between the fungal primers and other primers is that it contains inosine. We currently think that maybe the uracil-DNA glycosylase (present in the qPCR master mix but not the PCR mix, and also not when the primers have been used for qPCR in other papers) accidentally interferes with the inosine? This doesn't make much sense and we haven't found anything to support it (or refute it).
Does anyone have any ideas on what would cause a primer set to amplify with PCR but not at all with qPCR?
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Hi Maggie!
I have the same problem with bacteria (Pseudomonas).
I tried to change several parameters, but have not had any success yet.
Did you find any solution?
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I heared, not all Taq polymerases work well with DNA from environmental samples (water). We are using Promega GoTaq G2 Flexi and it does not work.
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There is no answer to this question. Different environmental samples can have different inhibitors which might interrupt the functioning of certain polymerases. So, the PCR reaction need to be optimized for the sample type and polymerase type.
Although, there are many polymerases available in market which claims to be more robust, still their efficiency differ based on the sample type.
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I would like to determine the functional role of a non cultured or eDNA based given bacterial community. Is there any online tool available which could determine the possible functional role of a microbes or its community using of 16s V3 / V4 region gene sequence ?
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For bioinformatics analysis with 16S amplicon sequencing data, this article may help you. https://www.cd-genomics.com/bioinformatics-analysis-of-16s-rrna-amplicon-sequencing.html
There are currently three powerful tools for predictive metagenomics profiling (PMP): PICRUSt, Tax4Fun, and Piphillin.
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I wanna extract eDNA from fish liver and blood, faced problems in extraction process. Please suggest valuable information.
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18 answers
Hello!
I'm trying to perform a TaqMan qPCR to check for certain fish species in environmental DNA samples. I am using a published primer + probe set. The probe is carrying FAM at it's 5' end and MGB-Q500 at its 3' end. When running the qPCR, I see no increase in the fluorescence signal. However, when I run the reaction on a gel, I do see nice PCR products of the expected size (same when I run a "conventional" PCR just with the primer pair), so I am assuming that my probe is either not annealing or something with the fluorescence signal detection is not working. I was running another TaqMan essay with similar conditions (same modification, same chemistry) on the same machine in parallel, which works fine.
Is there a way to test if the probe is annealing? I was thinking about running a PCR just with the probe as a substitute for the forward primer, but the 3' modifications would inhibit the extention anyay, right?
I'm using the Thermo Fisher Environmentam Master Mix 2.0 with the its standard cycling condions (which are also the conditions published)
Tms:
(calculated with Thermo Fisher Mutliple ologo analyser):
F-primer: 62,8°C
R-Primer: 67°C
Probe (without MGB): 63,6°C
Any help is very much appreciated! Thank you!!
Tamara
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To add: There could also be a problem with the machine's optics / detection if you can find the appropriate sized product when you run out the sample on a gel but the machine shows no amp.
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7 answers
Is Environmental DNA representing a significant trail in ecological studies? Rather than the traditional methods in ecological survey particularly in aquatic and soil studies. Please we need your opinion on this topic.
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Environmental conditions affect significantly DNA fingerprinting. allele genotype frequencies were highly correlated with the envirment.
study focused on traditional methods in ecological survey can be confirmed with moilecular result. This can be a way for solid conclusions
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How to interpret each result of my electrophoresis visualization by gel doc?
I have 11 samples. Wells those marked by number 1 to 7 are DNA samples from alligator gar caudal fin, and wells those marked by 1L, 1N, 3L and 3N are eDNA samples from water. I think I did any mistake in my extraction techniques for my eDNA samples, so the results are clean. But how to interpret the others?
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My PhD student has been working with eDNA samples working on 2 genes namely COI (Leray's-313bp) and the MiFish-12S primers. He is using Illumina overhanging adapter with specific primers to amplify each samples, which means the primers became quite long with additional 30+ bp of the overhanging adapter (for each ends). He is now having problem with COI amplification. He did try a lot of protocols including Leray's touchdown PCR method, from using normal taq polymerase to using HiFi proofreading polymerase. And finally when he used HiFi polymerase there are some clear bands of non-specific amplification at 100bp+ region. However, we suspect that the bands are not the target size. With the COI primer, he is supposed to get 313bp, and with the addition of the overhanging adapters (+67bp), he is supposed to get a product with 380bp. But the product size is at 450bp+.
We need some advice here, because he is using pre-stained GelRed, and it may cause some issues with slow DNA migration in gels. Or probably, he did not amplified the right mtDNA. His positive control, labelled FISH did amplify at 400bp-ish. He repeated with the gel, but he still get similar band patterns. Is it possible that the target size is higher with the primer his using? He also gets a faint band on the negative control. He is sure that all his chemicals are fresh and he used fresh nuclease free water in the experiment.
The details of primers used-
mlCOIintF: 5'TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGGGWACWGGWTGAACWGTWTAYCCYCC
jgHCO2198: 5'GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGTAIACYTCIGGRTGICCRAARAAYCA
*the underlined nucleotides is the Illumina adapter
from: Leray, Matthieu, et al. "A new versatile primer set targeting a short fragment of the mitochondrial COI region for metabarcoding metazoan diversity: application for characterizing coral reef fish gut contents." Frontiers in zoology 10 (2013): 34.
Is it possible that the HiFi polymerase caused this 'contamination' to occur, because he did not see any bands or smearing for the negative control when he used the normal taq KAPA HiFi HotStart ReadyMix (2X) and buffer.
Can someone help us. We are at our wits end trying to figure out these problems. Thank you.
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I have not used the CO1 primers. For universal primers, it is normal that there is length variation among species. Mostly, you got targeted amplification based on your gel image. I would use DNA extracted from tissues of species known to be present in your eDNA samples to verify this.
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Dear Researchers
I hope you and your research is well. My own research is a little poorly. My Professor suggested I ask independent researchers about my lab-related problems. If you could answer the following I would be truly grateful.
I am developing a method to use environmental DNA (eDNA) to detect an aquatic macrophyte in lochs.
In the PCR lab, tissue samples were extracted, with a step that included the grinding of dried material in an open Eppendorf tube using a micro pestle.
During primer design, primers were tested using standard PCR. After amplification, the plates were unsealed, and processed using ZAG capillary electrophoresis to determine nature of amplicons of target and non-target species.
Once primers had been shown to be species specific, a probe was purchased and field samples were processed using qPCR (MGB-Probe with FAM Dye, and Environmental Mastermix). All samples, and many Non Template Controls (NTCs), showed contamination.
The samples had been extracted in a room adjoining the same lab where PCR amplification occurred, after primer design. Also, all pre-PCR work (preparing qPCR plates) occurs in this side room. A sliding glass door with a 20 mm gap separates the two rooms. 20% bleach was used for decontaminating surfaces.
I hypothesised that DNA contamination was from the air. In 2 repeated experiments that exposed NTCs (no template) to the air for different time periods. In both repeated experiments, I found that no contamination (0/12) occurred when wells were sealed immediately, but 5/12 replicates showed contamination when left exposed to the air for 30 minutes. See attached picture.
Questions
1. From my NTC experiment's results, can it safely be concluded that the air is the source of contamination?
2. What lab process or experimenter behaviour produced the contamination?
3. Could contamination be minimised by using qPCR and SYBR-Green Dyes to develop primers (e.g. you don't have to uncover the post-qPCR wells)?
4. If a hood with UV lighting had been installed in the adjoining room where sample extraction and plate preparation occurred, could this have prevented/minimised the contamination?
5. What steps, protocols and equipment is used in your eDNA lab to minimise contamination?
Many thanks
Nick Crutchley
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It's generally a bad idea to leave open your qPCR plates for any length of time, and definitely not 30 minutes (though I realise that was probably an extreme for the experiment). I must confess though, I've prepared a NTC mix, transferred to three wells with the same tip and two were clean, and one had a product in the melt curve - though it was obviously not my target. Is the peak the same as your product, and does it come out at the same time?
In general, the most likely sources of contamination are your pipettes and reagents. That would potentially explain your 5 false-positives. Having filter tips and fresh reagents (or jealously guarded reagents that only you use) would address these issues.
You may want to consider investing in a tissue milling apparatus (TissueLyser or similar), so that you can grind up your samples in a closed test tube.
Further, you could do the milling/extracting in a fume hood and the qPCR setup in a laminar flow.
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I will need to identify the species of bird from faecal samples using metabarcoding. Are there any primers that are short enough to do the job? The traditional COI primers are too large.
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agree with@ Gabriel Gonzalez
regards
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Looking expert opinion...
I have collected marine sponge samples and were shadow dried for two weeks. Now the sponge samples are well dried and can be directly powdered by grinding. I would like to study the sponge associated actinobacterial populations (uncultured) from the dried sample rather than fresh sample.
Here comes my doubt,
If we grind and use the sponge powder for metagenomic DNA extraction, does the DNA be damaged/sheared ?
or
can directly use the dried sponge material (without grinding) for metagenomic DNA extraction?
Kindly, some one clarify my doubts.
Thanks in Advance,
Siva
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Can't say much about the effect on the microbial community in the dried samples, but I feel using dried samples directly would be a better option if we consider the bias introduced during the sample preparation. Rule of thumb, more the steps involved in the sample prep, more the bias.
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Hello, I want to transport environmental DNA samples obtained by water filtration. They are therefore packaged in filter cartridges (envirocheck HV) containing a storage buffer. I wonder about the potential impact of X-rays (during airport baggage checks) on the integrity of DNA molecules (shearing and modifications). Is it necessary to take special precautions? Thanks Cédric
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I'll like to applied eDNA in water quality of Tigris river.
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I follow
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Hi,
I'm doing metabarcoding analyses with eDNA samples and after clustering my sequences (97 % similarity) and did a blast (97 %) with a reference data base (UNITE), I notice that some OTUs have the same species identification. Logically they should have been grouped together during the clustering step.
Thank for your help
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Hi Genevieve, I agree with the answer above.
Your sequences can be >3% different to each other and therefore clustered in separate OTUs. However, when these OTUs are blasted, a taxonomic assignment is made based on sequences available on the dataset you are using (e.g. GreenGenes; NCBI ). Sometimes the taxonomic assignment is "forced" to the closest match (which, paradoxically, could be only 80-85% similarity). This is why your OTUs, while >3% divergent between each other, are assigned the exact same "taxonomical ranking".
How to deal with this (in my opinion):
Always remember to check the matching % of your OTU with the assigned taxonomy. If necessary, manually blast it yourself and check how similar is the most similar sequence. If your OTU is only 80-90% similar to the most similar sequence available on the dataset, this should be what you have to report and to keep in mind.
For example, an OTU 85% similar to a (e.g.) Sodalis glossinidius sequence is NOT a Sodalis glossinidius sequence. It probably belong to the same genus or family, but it would be a big mistake to report it as that species..
Hope this help!
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We need to clone environmental DNA (eDNA) into heterologous expression hosts in order to produce natural products but this eDNA is such a huge piece of DNA (20 to 150 kDa) that we need to construct a metagenomic library first and then make a PCR-targeted sequencing to find parts of Biosynthetic Gene Clusters (BGCs) and, finally, joint the pieces together into a BAC (Bacterial Artificial Chromosome)
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Shotgun sequence can be a better solution. In these case Ilumina or PackBio are the option to choice, other platform also there, but previously mentioned will be better . Felipe, thanks and regards.
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Most published work deals with freshwater samples, and one specifically concludes that using lysis buffer or samples dried with silica bids yield better results than preserving in ethanol. Would this conclusion be the same when dealing with saltwater samples? or would the different chemistry composition of saltwater yield different results for preservation?
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It must be same. However, thorough literature survey is required before ascertaining.
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I will be collecting water for microbiome and eDNA work and I am trying to figure out the best methods for cleaning/sterilizing my plastics.
Water will be collected using plastic turkey basters and filtered with a metal strainer into a plastic container ~2L capacity. For each sample, I will be using a different baster, strainer and container that will be brought to the field site sealed, but I need to be able to clean/sterilize these items in between sample visits.
With arthropod surface sterilization, I've used cleaning with bleach, sterile water and ethanol. A similar set up might work for the plastics, or maybe a combination of cleaning with each of the above and then placing under UV. If anyone has experience with this type of cleaning or any suggestions, I would greatly appreciate the input.
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Thank you for this info! I have typically seen 20% bleach used, I wasn't aware that 1% would be sufficient. Does 1% work to degrade the DNA (so that it won't be picked up when PCR amplified) or only to kill whatever bacteria or fungi might be present on the plastics?
Also, do you do a wash (water or ethanol) after the bleach?
Thank you again for your input.
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Saline temporay lake. Fixed with lugol.
Someone can help me with the taxonomy
thanks in advance
Maria 
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Thank you so much for your answer Bohuslav Uher, all information helps.
My Gymnodiniums are biger: 18-19 x 17 micrometer more o less, i was thinking  perhaps in Gymnodinium cnecoides
Jiri Popovsky and Lois Ann Pfiester (1990): Dinophyceae (Dinoflagellida) . Gustav Fischer Verlag. Stuttgart. p. 103-104
Best wishes
Maria 
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Hii
What is the source of extracellular DNA (eDNA) released from viable bacterial cell independent of cell lysis and bacterial cell death.How eDNA produced and eject from the cells.
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QIAamp stool kit (51604) worked well but QIAGEN didn't produce anymore and they have a new fast stool kit (51604) but this new kit didn't work. Any suggestion or modification to do the isolation from fish feces marine fish feces (white fish, rainbow troat, and salmon) would be helpful.
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Sheila, hi.. I think I misunderstood your previous question.
maybe you can still use the fast stool DNA kit but you just need to do a pre-treatment before the DNA isolation from the parasites... I thought your samples are from stool/excrement! You can check several options of pre-treatment method from the Qiagen book/catalog...depending on your sample conditions!
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Hello all,
I'm currently working on a project involving environmental DNA. I'm at the PCR step and can't quite figure out the appropriate amount of DNA template to use per 50 micro molar reaction. 
I do not know the concentration of my DNA since it came from samples of water I collected from streams. 
 I'm using a primer mix of 100 micro molar concentration, each primer in the mix is at a 2 micro molar concentration. 
What is an appropriate amount of DNA template (in microliters)  to use per reaction? 
Thanks in advance. 
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Is your protocol comparable to any published study on eDNA, particularly one used for similar taxa and a similar goal (detection only, enrichment for NGS, etc). I can't tell what kind of eDNA you're talking about (relatively abundant microbial or rare macrobial), but if it is macrobial aqueous eDNA, you might consider these recent papers that provide some guidance on protocol design for species detection:
Furlan EM, Gleeson D, Hardy CM, Duncan RP. A framework for estimating the sensitivity of eDNA surveys. Mol Ecol Resour. 2015; doi:10.1111/1755-0998.12483
Schultz MT, Lance RF. Modeling the Sensitivity of Field Surveys for Detection of Environmental DNA (eDNA). PLoS One. 2015;10: e0141503. doi:10.1371/journal.pone.0141503
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I am working with degraded eDNA (bird poop, dead insect larvae, etc.) and I usually use BSA but this is the first time I will be using a ready-to-go PCR mix. I was wondering if such mix is already optimized for degraded template DNA so that I wouldn't need to add the BSA anymore. Thanks!
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thanks pierre!
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I will be collecting water samples for an eDNA study on Hellbenders and would like to test the water in the streams to figure out if endocrine disruptors are leading to their decline. Is there a solid test that exists to test water samples for estrogen? That can be done by a graduate student? Thanks!
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I don't have experience with this specific product, but there are ELISA test kits for detecting estrogenic hormones in water - http://www.biosense.com/render.asp?segment=3&ID=87
Other ELISA tests I have been involved with were relatively inexpensive, and appropriate for a grad student with lab experience.
I've seen one stat that claims 80% of US waters have detectable levels of estrogenics present, and that they're even in bottled waters. So don't be surprised by positive results. So, one problem for you to address is correlation versus causation.
Cheers,
Mike
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I have barcoded 18S and 16S rRNA amplicon libraries from 25 marine environmental DNA sampling stations, for a total of 50 samples. I would like to sequence these on an Illumina Miseq, but I will need to pool these libraries myself prior to submission. I have never performed pooling myself, so I had a few questions about this!
The submission requirements at the facility I plan to use call for >10nM of DNA in more than 15uL of buffer.
My question is, what kind of volumes should I work with when diluting and pooling my libraries to equimolar concentrations? Also, for the number of indexed samples that I plan to sequence, what should my final library volume approximately be? My PCR product was eluted with Qiagen EB buffer for a final total volume of 30uL.
An Illumina protocol I looked up recommended 5uL of diluted DNA per sample in the final library, but this suggests that the final volume submitted for sequencing could vary wildly depending on your number of samples, which seems odd to me.
Thanks in advance!
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Solomon is right.  You just need to be over the required amount, as your sequencing core can always dilute down.  The first step in a sequencing run is clustering, which is spreading the fragments across the Illumina chip such that they are not too concentrated which results in the inability to correctly sequence nor are they spread too thin which is a waste of sequencing reagents and results in a low yield of sequences.  
Also, how are you quantifying your library.  Be careful NOT to use a straight Bioanalyzer, Nanodrop, Qiaexcel, Qiaexpert, or Qubit result.  These all tell you your nucleic acid concentration, which you will still not know your 'sequence-ready' library, which contains the necessary adapters for adhering to the flow cell.
Use the Quantimize kit to measure the precise concentration of sequence-ready library as well as QC:
Otherwise, you will be wasting your sequencing
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Dear all,
One of my friends has to amplify some species-specific genes from cooking oil, which contained little and mostly degraded DNA. I think paleontologists often work on degraded ancient DNA. Could someone suggest me a protocol? My friend has isolated the DNA using a membrane-based method. Now it's all about the primer design and PCR. Thanks in advance
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That seems a rather good choice.
There is nothing you can do regarding PCR buffers etc. Your only option is to select small size products and hope at least one fragment of your fragmented DNA contains the whole sequence.
Thus, you could also help by increasing DNA concentration (in the hope that such molecule containing the whole sequence gets into the reaction). However this only works to a certain limit.
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I'm planning to perform PCR amplification on environmental DNA (filtered microplankton biomass from seawater samples) for both 18S and 16S, using Taq Hifi polymerase and barcoded Ion Torrent primers.
My 18S forward primer Tms range from 72.1 o 72.6. The reverse primer is listed at 65.2 degrees.
My 16S forward primers are 73.6 to 74.8 degrees, and the reverse primer has a Tm of 67.3 degrees.
What I'm more curious about are the optimal lengths for the denaturing, annealing, and extension steps. I am planning to test how well my samples amplify, perhaps using a temperature gradient, but I'm wondering what kind of run settings you might suggest for a starting point!
Similarly, what master mix/primer/enzyme/template mixture might be a good starting point? My primers are diluted to 10 uM.
We do not typically use Ion Torrent, and I've never performed PCR before using Ion Torrent primers, so I'd appreciate it if someone could point me in the right direction!
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Just start with optimal annealing temperature for your polymerase enzyme and your primers. Perhaps use a gradient PCR to minimise primer-dimers and wrong targets.
I have successfully used Bioline MyTaq Hotstart ready mix, KAPA Robust ready mix and KAPA HiFi HS Readymix for Ion Torrent.
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Ranges I have seen are between 30bp-80bp. 
Are there chances fragments could be longer? Especially ones recently released into the environment?
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I think you will have any length of DNA in the water, but as smaller the fragment get you will catch more species. So it is mostly a compromise between how long it should be to answer my question but get a maximum of species. Theseis always depending on your question as well as the ecosystem, leads to the normal final conflict of Biodiversity against specifity.
I working in marine water with 120-170 bp and try to be less than 200bp and I have good experience with that. So it should also work for aquatic system, mostly even better. But the problem is more that DNA degenerated differently, even within mitochondrial DNA I suggest large differences between regions based on my results.  
 Hope these give any helpful information to you!
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I'm hoping to compare field sampling data with the results of fish species determined to be present using  eDNA metabarcoding. Will likely send out for the high-throughput portion, but everything else will be done in-house.
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From experience –attempting to do this for stream inverts– nothing in the world of eDNA is straightforward!  I'm probably telling you what you already know, but any semi-quantitative e-system that is going to be sufficiently accurate for reliable use requires the testing of many potential targets and many mixtures of different compositions before you can ever get near using the system in the field.
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I am currently running an experiment trying to detect environmental DNA for the species Hemimysis anomala or commonly known as bloody red shrimp.However, while running my controls using actually bloody red shrimp tissue, my gels were negative. I am following the protocols and using primers based off of two papers: 
Audzijonyte, Asta, Karl J. Wittmann, and Risto Väinölä. "Tracing recent invasions of the Ponto‐Caspian mysid shrimp Hemimysis anomala across Europe and to North America with mitochondrial DNA." Diversity and Distributions 14.2 (2008): 179-186.
and
Questel, Jennifer M., et al. "New data on mitochondrial diversity and origin of Hemimysis anomala in the Laurentian Great Lakes." Journal of Great Lakes Research 38 (2012): 14-18.
I am using Qiagen Blood and Tissue Kits for DNA extraction. Amplifications were performed in 20 uL volumes containing 0.5 U of Hot Start Taq DNA polymerase (Qiagen), 1 x buffer, 3 mM MgCLs, 0.2 mM dNTP, 10 pmoL of each primer and 10 ng of genomic DNA. DNA samples were run through a thermal cycler at conditions as follows: 95°C for activation for 15 minutes, followed by 35 cycles by 1 minute at 95°C, 1 min at 50°C, and 1 min at 72°C, with a final extension of 72°C for 5 minute.
PCR products were purified using QIAquick PCR cleanup kit and run through Agarose Gel Electrophoresis according to standard protocol. 
However, I am unable to detect any DNA and before I can begin my actual amplification of my eDNA filters, I need to be able to ensure that the primers and my technique can detect tissue of the H. anomala I am working with from the Finger Lakes region.
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when you say no measurable dna do you mean that you cannot see any bands after pcr. If so the problem should be isolated to dna preparation or improving the pcr process. to test the dna preparation borrow/make a primer set other than yours and check that they work on your dna. Similarly borrow some dna that works for other people and run your primer set on it. Often I find that when primers do not work it is because one or both primers have degraded due to being dissolved in water not TE or degraded by freeze thawing too often. Primers do not work also if the 3' end of the primer anneals over a polymorphism but if these are old inherited primers from another researcher buy them freshly made. Also check the sequence of your primers against the shrimp genome...it is not unknown for typological errors to creep into publications at the manuscript preparation stage particularly on primers which are not amenable to spellchecker
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Can anyone suggest how I might go about constructing power curves for permanova tests? I am working on an eDNA metabarcoding community impact study and want to investigate how many replicate subsamples should be analysed in future monitoring surveys.
Thanks in advance for your help!
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Do you have Minitab? Set up data file, do analysis, click stat, then power and sample size, choose type of anova, enter parameters, click okay, then you will have your power graph!
Andrew :-)
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Dear all,
I'm looking for detailed protocols for extracting DNA from soil, detect plant species in soil DNA samples, and quantifying relative frequency of species DNA.
I find some good articles, but I need detailed protocols.
Thanks.
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While not searching for plant DNA I also use the MoBio powersoil kit to extract community DNA for analysis. It gives very high DNA percentages around 90%+ is very easy to use and relatively inexpensive. In addition you can likely request a free sample from MoBio to test out to see if it works for you.
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I am experiencing a contamination problem in my DNA extracts. I did DNA extraction from water samples using phenol/chloroform. However, I do get a shoulder/high peak at 230 nm whereas there is a slight peak at 260 nm when measured with Nanodrop. My 230:260:280 ratio is more like this: 1.6:0.8:0.5.
I adapted my fieldwork sampling protocol from Foote et al. (2012). Based on the literature search, and similar problems people have experienced, I do think that the high peak at 230 nm is due to some salt contamination along with residual ethanol. I am in the midst of troubleshooting, but I am not sure how to fix this problem. There are people suggesting to use ether (to remove organic solvents) or to dialyse the probe if salt is the contaminant. Also, during PCR some additional reagents can be used, such as BSA and/or glycerol to prevent any potential inhibitors getting amplified.
Any suggestions on how to carry out? I would like to have purified DNA before troubleshooting with PCR step, ideally.
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Starting with the quickest and easiest things I can think of:
If it's residual ethanol, you can often just warm the samples in the water bath at 50C for 30 minutes or so, and the EtOH will evaporate.
If it's residual organics, a second extraction with straight chloroform will help remove them. 
Finally, a second precipitation can't hurt (add 1/40th volume of 5M NaCl and 2x volume EtOH, let precip for an hour, spin max speed for 10 minutes, then wash with cold 75% EtOH)  -- It can dilute out some other salts that come along for the ride.
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Hi all,
I was hoping that someone could tell me of a protocol to isolate DNA from soil samples collected at a mining site - preferably one that doesn't need a kit and could be achieved with a CTAB modified approach. The current techniques I'm using are more suited for humic soils (Verma and Satyanarayana, 2011) than for the gravelly heavy metal contaminated collected soil samples.
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Hello, 
My superviser provided me some plant samples but the samples are in very bad condition. I am using PVP and sodium bisulfite in extraction buffer for phenolic compounds and normally it works well. After extraction I did chloroform:isoamylalcohol cleaning twice. After isopropanol precipitation I got phenolic contamination. Has anyone ever tried coloumn purification of samples after manuel extraction?
If so, what should I do? I have Qiagen plant DNA kit and Thermo Plant DNA kit.
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Hi,
when we purify DNA or RNA from problematic samples, the column purification of the isolated nucleic acid is a routine method. You will of course loose some DNA, but that is usually not a problem, whereas purity is of main importance (if you have enough starting material).
I had such problem of extreme polyphenol and polysaccharide content of old grape leaves, so I could suggest you to do the next:
You can follow the RNA isolation protocol of Karen E Reid et al. (http://ccforum.com/1471-2229/6/27) up till the point of selective RNA precipitation, which step would then be replaced with NaOAc/EtOH or other kind of DNA precipitation, RNase treatment and/or another column-based DNA purification, whichever seems necessary with your samples.
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Dear Donald,
I have had same experience with this kit. I contacted  MO BIO Technical Support <technical@mobio.com>. Their reply is as follows:
"Clay soils usually give low DNA yields. We typically get yields below 10 ng/ul from high clay soils. There are some things you can do to maximize the yield. We have found that using 0.1 mm glass bead tubes along with a high powered bead-beater can boost the yields for this soil type. These bead tubes are in our PowerLyzer PowerSoil DNA Kit (12855). You can learn more about this from our Application Note. See the attached file. If you want to try the glass bead tubes, Research Instruments should have samples of this kit (12855-S).
f the soil is not too high in humics we have an alternative protocol for low biomass samples that uses phenol:chloroform:isoamyl (PCI) that and might also increase your yields. I'll copy it below. You could try this first. In addition to the PCI you would need 100% ethanol.
Even though you can't accurately measure the quantity on a the nanodrop you may still have plenty for PCR. 10 ng would be enough for 16S PCR.
Alternative PowerSoil Protocol for RNA and DNA from Low Biomass Soil
1. If starting with the dry glass bead tube from the PowerLyzer PowerSoil Kit, add 0.25 grams soil followed by 500 ul of Bead Solution and 200 ul of phenol:chloroform:isoamyl alcohol pH 7-8 (We buy Amresco brand, catalog# 0883- it comes with the buffer for the phenol). If using the original PowerSoil Kit, remove 200 ul of bead solution from the tube and add in 200 ul of PCI.
2. Add the 60 ul of Solution C1.
3. Vortex 10 minutes. Centrifuge to pellet (1 minute full speed).
4. Remove the supernatant or upper aqueous layer if you have one, to the new tube. Add Solution C2. For low humic soils, reduce it to 100 ul. Add 100 ul of Solution C3 next and mix, and incubate at 4C for 5 minutes or on ice.
5. Centrifuge to pellet (1 minute full speed) and remove the supernatant to a new tube.
6. Ideally you will have 650 ul of lysate and can add 650 ul of Solution C4 and 650 ul of 100% ethanol. If you have 700 ul, add 700 ul of Solution C4 and 600 ul of 100% ethanol.
7. Load the lysate 650 ul at a time and bind in three steps, alternating with centrifugation or using a vacuum manifold.
8. If the membrane is not stained brown, wash with 650 ul of 100% ethanol and then 500 ul of solution C5. If the membrane is stained, per sample, prepare a mix of 300 ul of Solution C4 and 370 ul of 100% ethanol. Wash the column with this mixture first. Follow this wash with the 100% ethanol and the Solution C5.
9. Dry the spin column for 2 minutes full speed. Transfer to a clean tube.
10. Elute in 60 ul of buffer C6. Let the buffer sit on the membrane 5 minutes before elution."
I wish it would be useful to you.
Thank you so much and wish you success with your research.
Pardis
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i have mixed bacterial DNA sample. to identify all bacterial community present in the DNA is DGGE technique is useful for identification of all bacterial community in DNA sample
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Yes you are right DGGE will be best for you. Another technique which you can use, is very similar to DGGE is Temperature gradient gel electrophoresis (TGGE).
You can also also go for clonning.
Amann, Rudolf I., Wolfgang Ludwig, and Karl-Heinz Schleifer. "Phylogenetic identification and in situ detection of individual microbial cells without cultivation." Microbiological reviews 59.1 (1995): 143-169.
Muyzer, Gerard, and Kornelia Smalla. "Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology." Antonie van Leeuwenhoek 73.1 (1998): 127-141.
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I need to store sediment samples for extraction of RNA to do metatranscriptomic analysis but the RNA extraction cannot be performed before around 6 months since sample collection and shipment. No problem with keeping at -20 or -80 °C, but is that enough or should I add RNA later or similar? I read that these chemicals don't work with sediment, is that true? any suggestions? Thanks!
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RNA extraction from the sediments will be frustrating anyway. If the sediments are rich in microorganisms then perhaps you will have less to worry about...
I wouldn't use RNAlater - it did not work for us at all when we tried to preserve RNA in mussel tissue samples; its use is tricky and does affect downstream isolation procedures. In our case simple 70% (final) ethanol worked the best. But in your case, if -80°C is not a problem I would say - go for it. Minus 20 is a little to high for long term storage of crude samples, you will almost inevitably have more degradation then. It would be advisable to actually run a series of extractions from the typical sample of yours conserved with different methods to see how they affect you chosen isolation procedure before wasting the whole sampling effort.
Good luck.
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I am working on a project trying to isolate  intracellular DNA (iDNA) from a soil sample. I have been trying different approaches to mute or block the amplification on extracellular DNA (eDNA), and I am now trying to wash the sample to free up any eDNA fragments that may be adsorb to soil particles before the treatments that I am applying. I have read one paper that used SDS at 0.1% concentration to (what I assume) wash off or desorb the DNA from the particles. I am still very new to molecular research and throwing soil chemistry on top of it is a little challenging, although exciting to learn. does this seam reasonable, using low conc. SDS as a detergent, or is there a better solvent?
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Hello! 
I suggest that first you need to use some cell wall degrading enzymes or treatment to allow SDS to show it best effect, by which ultimately you can recover the intracellular eDNA.
All the best.
Jagat 
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Hi,
When applying a Taqman probe (FAM dye & NFQ-MGB quencher) to a ten-fold serial dilution of tissue-derived DNA for eDNA quantification via standard curves analysis, I do not get the archetypal sigmoidal curves, but a series of increasingly more shallow curves that do not asymptote and seemingly would go on forever with unlimited reagents (see first fig). Compare the same reaction using SYBR, where we have the nice, beautiful sigmoidal curves (2nd fig). Both pictures are derived from reactions of 250nM F & R primers and the Taqman had 200nM probe. Varying either has no effect on the shape or decreasing pattern of the curves, but does change the height of them, viz. Taqman (I've also tried two Taqman mastermixes -  (Applied Biosystems Universal PCR Mastermix I and Mastermix II, both no UPG)). Both reactions used the same conditions (denaturation for 15 secs followed by annealing.extension of 1 min as per manuals, following enzyme activation step(s) for each mastermix).
I suspect that the probe is just inefficient, but given the limitations for unique primer-binding sites for this species - and therefore increased limitations for useful primer.probe pairs re: species-specific eDNA detection - I want to exhaust all possible options before reverting to SYBR green analysis, as Taqman is the gold-standard. Further, the probe is 27bp long (acceptable as far as I am aware), but some fellow in the lab suggested excising a few nucleotides from the 3' end - is that feasible?
Thanks.
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"as Taqman is the gold-standard" - who sais that?
Taqman has two advantages: it can more specifically detect (! not quantify!) the presence of a sequence in a mixture, and, therefore, can be used to multiplex.
But it also has disadvantages: there is no possibility to run melting curves, so the specificity of the amplification (not the signal generation!) has to be ckecked by old-fashioned gel electrophoresis.* The dyes sometimes show baseline-drifts, either possibly because of eximer-formation or because of quencher-loss.
For a quantification in a singleplex assay, Taqman probes do not have any advantage over SYBR Green, but it is usually a bit more expensive.
---
If unspecific products are ampified but not detected, their amplification can still compomise the amplification efficiency and so render the Ct values wrong. Not measuring/detectin a problem does not mean that the problem does not exist.
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I have several 1L bottles frozen at -80 deg.C and need to filter them for eDNA analyses. What is the safest way to melt the water?
i'm testing with two samples by leaving them at 4 deg. C but it takes a very long time (>1.5 days and its still half frozen) and i'm unsure that it won't damadge the DNA, since part of the samples is already liquid since approx. 1 day.
Thanks for any comments on this!
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Thank you for your comment and suggestion!
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I want to know which of these options apply during fieldwork.
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Hi Carlos: We do a lot of DNA extraction from plant tissue, so we dry it by using silica or freeze dryer, and its work very well.
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I have environmental DNA samples that were filtered through cellulose-nitrate paper and extracted using Phenol-Chloroform.
Samples were collected from small and ephemeral waters, and thus have high environmental contaminants (tannins, humics, etc.).
Tissue-derived DNA extracted with Phenol-Chloroform are not inhibited, so I suspect residual phenol is not a culprit. Besides, samples are washed with ethanol during the extraction after the phenol steps.
qPCR is conducted with SYBR Green.
What I have tried:
Dilution - I ran a dilution series. At 1:100, almost all samples are still inhibited. At 1:500, almost all samples are too low in concentration and fall under detection threshold. 1:200 seems a fair compromise, but some samples are still inhibited and some are too low concentration.
BSA/DMSO - Using both, samples are uninhibited in standard PCR. However, BSA seems to interfere with SYBR dye. DMSO alone does nothing.
Zymo One-Step PCR Inhibitor Removal Kit - this kit appears to have introduced inhibition to samples which were not previously inhibited (see attachments)!
Attachments (all samples are expected or confirmed positives, standards are green/yellow): 
DilAmpPlot (top) - samples diluted 1:200
DCCAmpPlot (middle) - samples processed using Zymo DNA Clean & Concentrator kit (does not remove inhibitors)
OSAmpPlot (bottom) - samples process using Zymo One-Step PCR Inhibitor Removal Kit
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If you still have dirty pellet and facing with inhibitors, I would offer you different protocol. An alternate and fast approach is using Norgen Biotek kits for environmental samples. I have dealt with plant-derived inhibitors and designed my own protocols to cope with them without using Qiagen products which were not efficient enough. 
You may download my book from RG and review the sample preparation methods for PCR in Chapter 2.
If you have further question, please get back to me.
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but some article say:Because of the low concentration of eDNA, special precautions must be taken for the amplification step. Firstly, the number of PCR cycles may be increased compared to traditional DNA work, to more than 50
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Hi Wenhao. There are many reasons to avoid setting more than 35/40 cycles for a PCR:
In particular, due to the exponential amplification reagents are used completeley at some point, dNTPs in particular.
The activity of the enzyme, despite being heat-stable is declining over time and in conclusion if you run the PCR for too long, you will get more and more side-products (mostly primer dimers, but mis-aligned primers can also make problems).
Those are the problems using a standard taq or master mix. Maybe if you use a specific kit for environmental DNA it is different.
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I will sequence DNA and RNA from grassland soils using Illumina to see differences in composition and functionality. I will use DNA sequencing as a reference to align RNA. I need to establish the paramenters to do the senquencing, 2x100? 2x140? 2x200? I also need to know the minimum reads per sample i will need to see results... 20M? 45M? 60M?
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OK, there are two possible concepts of functionality: one is for genes encoding enzymes acting in important metabolic steps, or another is about the genes that are actively transcribed vs the ones that are present and had been selected for a long time in the community. Is your analyses on total metagenomic/metatrasncriptomic datasets or is it on PCR amplicons targeting some genes in DNA and RNA (cDNA templates) environmental extracts or is one single organism? Depending of the complexity of the community (in soil is extremely diverse and a huge number of sequences will not cover the diversity of the metagenomes), then it is possible that you may run comparison of the reads that are found in both datasets. But this results will have always the limitation about how representative they are of the total RNA or DNA content. Is not an easy question to solve. As an starting point I would say, try to get the same amount of sequences on each dataset (metagenomic and metatranscriptomic to avoid normalizations and loosing information). And try to get as much as possible. As a guide, if you want to have a representative assembly of a genome, you need a minimum of 20X coverage with a 2x150 pair end library (read average 300 b). If an average bacterial genome is 4 Mbp, so you may need 80 Mpb of information for that single genome. Now, assuming that in soil community you have around 5000 different species, and 4 or 5 are above 1% in that community, to have a, let's say 20X representation of one single genome of one of the abundant species in the community, you may need a metagenomic dataset of 8000 Mbp (8Gbp, a full MiSeq flow cell). In the case of metatranscriptome is more difficult to do calculations as there the expression would be predominantly masked by rRNA over mRNA. And from the mRNA derived sequences (after RT PCR sequencing), can be classified with predicted function or mapped to the metagenomic dataset. 
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We are filtering river water, of different salinities. With some samples, even when spiked with DNA of the species in question, we get no amplification. We've tried environmental mastermix.
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Did you try to desalt your samples on spin columns?
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Just checking if I'm thinking about this the right way or missing something. I'm a lab newbie and haven't had to analyze samples before for use in whole genome sequencing. We are required to have at least 10 ug in the samples we send out for sequencing. Thanks!
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In addition what said above, make sure to use the diluting water and buffer to zero your instrument. Make sure to clean up the instrument with clean tissue and sterile water before each measurement. 
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I am looking for previous reports that showed sample variations of extracted DNA amount (and quality) from DBS card. There might be huge variations of DNA amount as well as quality of the DNA. This would be very important to know before we amplify target genes from the samples. 
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I don't have direct experience with this but this slide presentation could be helpful.
Thanks
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I am using morphology to identify morphospecies of freshwater invertebrates.  Some of their adult stages (when it comes to insects) have been sequenced using the 5'.  Apparently, it is 3' region that is most useful for NGS but I cant seem to find relevant literature explaining this.  
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If I analyze bacterial population of a  sample (soil) by 16s rDNA targeted Next Generation Sequencing (from community /environmental DNA ) then the method  should be called as Metagenomics or Metabarcoding? 
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No, it is not metagenomics. In the same way that you don't do genomics of an organism by studying a single gene. Think about it, you don't call genomics when you use 16S to identify a strain or describing a new species.
The Genomics Standards Consortium even manage the Minimum identification about X sequences with different standards, MIMS and MIMARKS for metagenomes and marker gene sequence studies respectively.
Appropriate terms would be 16S rRNA gene survey, amplicon sequencing, metabarcoding or even metagenetics.
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I need to do HLA Profiling "genotyping all types" for about 500-1000 samples. I'm willing to pay the Lab. fees if possible.
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Medical Research Institute, Alexandria university
Alhadara, Alhoria st., beside Almuasa Hospital
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We follow a protocol which is very similar with regular phenol chloroform method to isolate extracellular DNA from planctonic bacterial cells.
According to this protocol, we remove cells by centrifugation and filter supernatants. To degraded proteins and other components, filtered supernatant were mixed up with  phenol-chloroform–isoamyl alcohol. Then mixture was centrifuged. The aqueous phase was removed and mixed with sodium acetate and 100% 2-propanol. The mixture was then centrifuged, the supernatant was decanted, and the precipitated sample was air dried and suspended in dH2O.
After then, eDNA was measured by NanoDrop and results are between 2000-3000 ng/mL.
Agarose gel electroporation was used to visualize eDNA samples. But, in this proses no eDNA samples were observed.
What should we do? Do you have different protocol? 
How can we observe eDNA on agarose gel?
Thanks in advance!
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You must check your buffer, protocol for agarose gel. This might be the reason. As soon as there is a reading for DNA in NanoDrop, that means the problem was in agarose gel 
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If anyone can point me towards references, it would be greatly appreciated.
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Dear Kathryn,
You may find the following pre-print of our group useful: https://peerj.com/preprints/2044/
The BF2+BR2 are great, or the BF1+BR2 in combination.
You can also use the new PrimerMiner software (http://onlinelibrary.wiley.com/doi/10.1111/2041-210X.12687/abstract?wol1URL=/doi/10.1111/2041-210X.12687/abstract&identityKey=e14200f3-8855-4698-94a3-ad73c1b2b217) to specifically check for primers matching your target taxa (if known). 
The primers from the papers above may not be optimal (the first ref is for vertebrates, the second is from our paper on amphipods and primers may not be degenerated enough).
Best wishes
florian
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I am looking for contacts of suppliers of Taq Polymerase (and associated reagents)  that is stable enough for Storage at Room temperature or at -4 °C for a few days. Any contacts will be appreciated immensely!
Aman
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Thanks a heap Sammy. Do you have their contacts or website details?  I now got Genscript only.
Aman
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I isolated DNA from couple of different cell lines. I obtained desired A260/A280 values. However I obtained low A260/A230 values showing the organic contamination of the DNA sample, i.e. salt contamination. Would that interfere with bisulfite conversion of the DNA?
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hello 
i do not think it will effect the conversion by bisulfite , but it can inhibit PCR.
regards . 
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I intend to detect the insect species that fed on leaves collected from the field and showing signs of herbivory by insects (but no visible insect remains on the leaves).
The DNA left by the insect, if any, is likely to be highly degraded (at least that is what I expect).
What extraction kit, primers, amplification method and sequencing approaches do you recommend?
Thanks! 
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If you do not know the species of insects that you are looking for, a DNA extraction is not going to solve nothing, because DNA sequeincing  have only been made for very few groups of insects. If DNA is extracted from a species that has not been sequenced, most likely you will not be able to identify correctly the species or group of insects. What you need to do is to sample insects and have them identified with a specialist taxonomist.
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I know of three marine mammal eDNA publications: Foote et al. 2012 (PLOS one), Ma et al. 2016 (Cons Gen Res), and Port et al. 2016 (Molec Biol), and a couple of other researchers (Scott Baker, OSU and Kim Parsons (AFSC, NMFS) who are investigating the technology but haven't published yet.  Referrals to anyone in the field would be most appreciated.  Thanks, Dennis  
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The USGS Wetland and Aquatic Research Center is conducting eDNA detection analyses from water samples for manatees.  Contact Dr. Margaret Hunter (http://edna.fisheries.org/category/mammal/west-indian-manatee-trichechus-manatus/) for details.  
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I am into SELEX to get aptamers for my target. I have been doing eDNA-protein interaction studies with gamma-p32 labelled isotopes to my eDNA.
When i try to estimate the amount of labelled eDNA as compared to labelled Library in a scintillation counter, cpm values derived from the instument sometimes are very weird. Visually on a gel i can see that library has more labelled DNA than my test eDNA's. But cpm values say the opposite.
As we need to normalize the labelled content for the assay, if i normalize based on the cpm values, i do not get any binding on my filters! It seems very weird.
Procedure I follow :
Label the eDNA
CPM counts taken
DNA content normalized based on the CPM values
Heat and cool eDNA's and Library
Prepare protein concentrations on 96W format
Mix eDNA and protein for 30min at 25C
Take 2uL for 0 time(input)
Add Zorbax beads to the mixture
Pass through the filter (vacuum)
Incubate the filter plate in -20c O/N
Read for radioactivity.
Any suggestions and downloadable papers or books are welcome regarding filter binding assays.
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Dear Chandan
I recommend you make them filter binding assays in membranes of nitrocelulose in a dot-blot system. This method allows you to use higher volumes of incubation and concentrate each sample. Uses samples with 10 ug of target protein as maximun. Perform the binding assays in solution and then filter it in the membrane from nitrocellulose, rinse three times with PBS. The membrane can be exposed it wet with folex paper and you can quantify the retained aptamer with autorradiografic films and later retrieve ligands concentrates in the dot. You can retrive them by heating to 95 ° C for 5 min at H2O miliQ and verify their presence and quantity by Cherencoff and later, the use aliquots for more amplifications (Radrizzani el. al, 1999 and Moncalero et. Al, 2011). Good luck,
Martin
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I have read that qPCR will be useful for the quantification step, but I am unsure if I also need to perform a pilot study to see how much DNA is shed from my target species to use this as a reference. Is this necessary? Finally, how do you go about attempting this when your target fish is elusive and protected from fishing -- and will be very difficult to house in a tank.
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Factors that affect shedding include time, temp, noise, pH, etc - the list is endless however some excellent publications are available...
http://pubs.acs.org/doi/abs/10.1021/acs.est.6b03114?journalCode=esthag 'Quantification of Environmental DNA (eDNA) Shedding and Decay Rates for Three Marine Fish' and http://onlinelibrary.wiley.com/doi/10.1111/2041-210X.12595/full 'Critical considerations for the application of environmental DNA methods to detect aquatic species' are both excellent detailed papers with full reference sections. http://www.sciencedirect.com/science/article/pii/S0006320714004443 ' Environmental DNA – An emerging tool in conservation for monitoring past and present biodiversity' covers the potential confounding factors including shedding rate
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I'm looking at using meiofauna community data from eDNA analysis for benthic environmental monitoring.  I would like to filter the eDNA dataset to exclude non-benthic taxa but am struggling to find a source with this information.
Can anyone suggest a database or publication detailing meiobenthic taxonomic groups which I could use to filter the eDNA data.
Thanks
Paul 
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I know that biofilms are really complex and that they are often visualized with GFP markers in bacteria itself. However, I was wondering if there is a specific marker, an antibody or GFP-tagged gene, to be able to call something truely a biofilm. Some sort of switch between planktonic/biofilm bacteria.
Is eDNA staining considered a good marker? And can this best be stained with propidium iodide or are there better ways?
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Hello, depends which biofilm you have, biofilm can be single or mixed-strain biofilms. depends from it you can have specific marker, an antibody or GFP-tagged gene for exact bacterial strain from which biofilm formed.
The DNA staining with DAPI which preferable stains dsDNA can be considered a good marker too.
Cell numbers in the biofilms can be determined non-invasively by collecting the total fluorescent density in an image, then dividing by the average fluorescent density of a single cell determined earlier in an experiment.
Good luck!