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I have a set of metagenomic sequence data from aquatic eDNA samples, and have been able to analyze the the bacterial/16S aspects of the samples but the program I use cannot analyze eukaryotic data. Does anyone have recommendations for programs that can be used to analyze eukaryotic metagenomic data?
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As an option you can do it manually by BLASTing your reads against 18s sets like SILVA ones. Next counting the relative presence of the species by assessing number of reads (and/or proportion of reads) aligned to a specific organism.
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When we isolate the edna from water sample generally 2-5 ltr water filtered and dna is extracted but in my case the concentration of dna is very low (7-8ng/ul) and there fore I am having problem in the amplification in pcr. So what is the minimum quantity required to check the amplification using qPCR mechine?
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Thank you @Zulaykha Khurshid for the replying
But the problem I am facing is that I can amplify my gene of interest with semi quantitative pcr with 20 ng/ul but if I use this concentration in real time the gene of interest shows undetermined
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I extract environmental DNA from a soil sample and sequenced using Illumina's next-generation sequencing technology.
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DEAR Amsale Melkamu
Genovo: De Novo Assembly For Metagenomes, Journal of Computational Biology: a Journal of Computational Molecular Cell Biology 18(3):429DOI:10.1089/CMB.2010.0244.
Meta-IDBA: A de Novo assembler for metagenomic data,DOI:10.1093/bioinformatics/btr216
GOOD LUCK
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Dear Ecologist friends who have worked with water or soil eDNA and understand the fundamentals of field data collection, processing and analysis.
I have a few terrestrial species that I would like to target with an occupancy approach, each of them travel through (in transit) water sources, and some are burrowing species. I have a few questions, and please forgive my ignorance I know very little about eDNA other than the core concept.
1. Would eDNA from water samples be an effective means to detect terrestrial species that only transit through water sources?
2. Would soil eDNA be an effective means to detect species that burrow? (i.e. taking the sample directly from a burrow site and BLASTing the metabarcode to match with the target species)?
3. What is the average decomposition rate of eDNA in water and soil? I mean how long could a species presence be detected in water and soil after they have been absent from a given area?
Any discussion is welcome!!
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Dear Russell
It's really interesting what you are working in..
I can give you some answers based on our field works..
1. Yes! The release of eDNA is an instantaneous process. We used to quantify, by QPCR, eDNA traces after few seconds of species placing in the aquaria (Yet, I don't know specifically about the situation in streams)..
2. Yes, also..We have preliminary data about eDNA traces from soil burrows made by the redswamp crayfish Procambarus clarkii, which invaded the Nile systen in Egypt. Being the burrows wet or dry, we could detect the species passage there, in its burrows I mean..
3. Don't know exactly the rates of eDNA depletion. I will search for it,that's an excellent concern..
All the best and good luck..
Khaled
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I work on environmental DNA and have really murky/sediment-laden water that I need to filter. The idea was to remove some of the sediment by centrifugation but I'm afraid that the DNA in the water would also be removed. I've tried to find articles or even similar questions but have found nothing (maybe i'm not searching the right key words?). I was wondering if anyone else had a similar issue or even knew where to look for an answer for this?
Thanks in advance!
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The DNA itself dissolved in the water well, the centrifuge cannot separate it from water. but the DNA in the bacterial, etc. will be separated by the centrifuge.
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I need the information about the companies in India offering services of environmental DNA for any organism. Please add website links.
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Hi Dinakarsami, I'd be happy to help you with metagenomics and metabarcoding analysis. Pl. drop your message.
Thiru
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I am interested in knowing if there are any eDNA field extraction kits - apart from the Biomeme field kit.
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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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Yes, I imagine that could certainly be a concern. For me, I was more interested in qPCR as being a more sensitive method of detection, but less interested in quantitation. I categorized samples presence-absence, based on critical threshold (which seems intrinsically subjective). With dilution, I had to accept the possibility of a few false negatives, but those would be in my most unreliable samples, anyway. If you are interested in absolute quantities, especially on the lower end of the spectrum, I'm not sure how to reckon with your concern, which I believe to be valid. I never had to consider those issues in my work. I know there were a handful of studies in the time I was doing my research that dealt in quantities, and I imagine there have been many more since. I don't know if any also dealt in PCR inhibition, so I wouldn't know of any good leads to point you towards.
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I'm looking to automate DNA extractions and I'm trying to figure out which option is better: getting a QIACube to use the DNEasy kits (which seem to have the highest eDNA extraction success of all the kits) or a magnetic bead based one like the Maxwell. I've seen one paper (Sanches & Schreier 2020) that compares beads & Qiagen DNeasy and says the DNEasy kits perform better. Have others had success using magnetic bead extractions on eDNA samples?
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Just wondering if someone has done it? We usually use environmental DNA (eDNA) for library preparation and I am wondering if eDNA concentration is very low, can we initially amplify the 16S region (1600 bp) via 27F and 1492R primers and then use the PCR prodcut for library preparation. Would be glad to hear from your experiences.
Thanks
PS: the concentration of eDNA is low and doesn't match the quality control criteria for a private sequencing company.
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Loos like you are trying to identify your bug based on full length sequence by using Sanger sequencing. In this case, yes you can re-amplify the amplicons from PCR product.
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I am planning to prepare a set of environmental DNA samples for WGS, in order to obtain functional information. I do not aim to get MAGs. But I realized that the quality of my original DNA material is not of so good quality. The concentration is so low, and the absorbance ratios cannot be properly assessed by using Nanodrop. Additionally, gel electrophoresis show a kind of smear.
My question is... is it WGS worth in this case? or should I forget about it?
Thank you
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Thr first question should be do I need a WGS?
Since you want to get functional information, WGS will not provide you any.
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I am looking for recommendations on purchasing a new qPCR machine. Are there any specific makes/models that perform better? Also, are there any special options that should be warrant special consideration? It will be used for several applications (eDNA amplification and expression profiles). Any suggestions would be appreciated.
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Try to used Machine learning
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I'd like to have your opinion on this subject. When publishing a paper or writing a report or thesis, if I refer to the DNA extracted from soil samples taken from a mesocosm experiment (i.e. containers filled with soil undergoing different hydrocarbon bioremediation treatments, kept outdoors at regular environmental conditions on the site of study, and sampled once a week during three months), should I say I extracted and sequenced to study the bacterial communities the total DNA from soil or that I extracted and sequenced the eDNA? Which one is the proper way to refer to it?
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Thank you very much for the link!! and so... if I use i.e. the mobio (qiagen) power soil dna extraction kit, I'd refer to microbial eDNA? or if I use a protocol using lysozyme, proteinase K, phenol extraction, ethanol precipitation to extract dna from soil samples to do bacterial 16S amplicon sequencing for example, should I mention microbial eDNA or just eDNA? what is your opinion on this?
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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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Hi Sivasankar Palaniappan ! How did the shadow drying method affect the DNA quality of your sponge sample?
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The parameters of the curve all seem acceptable, typically reactions results are above 0.9852, -3.1 to -3.6 slope, and 90-110% reaction efficiency. The results seem to be what I would expect from eDNA and the dilution series seems to be working well. But the lack of a plateau is making me doubt the validity of the results. Do you think this is an issue. What could cause it?
I am using primers designed for another project, temperatures that multiple authors have tested with them and confirmed, and reagents otherwise identical to their experiments. Increasing the number of cycles has no effect, and I have tried experimenting with reagent concentrations and total reaction volume without any luck. I am considering changing mastermix next, which will be pricey...
Thanks for the help,
Sam
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Thanks for the answers folks. The machine is an Agilent Aria MX. I believe it does have ROX correction.
I'll look into the other alternatives, re-run some qPCR's to check the melt curve, and perhaps do some regular PCR's to test a range of temperatures with my primers. Will get back to you.
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I have a large volume of P. leniusculus presence/absence surveys of small sites on up to 20 different rivers this year and it seems the jury's out on how accurate eDNA sampling methods are, especially in rivers. Any advice on the best approaches and how to deal with potential false negatives would be greatly appreciated.
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Nicola Green for your kind perusal
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I'm running eDNA samples that have super low DNA concentrations. When I prepare them for qPCR I would like to spin them down, and I have a small benchtop plate spinner centrifuge, but I'm not sure that I trust it to not mix samples when I place the plate in vertically with a plate film on top.
I see amplification at very high Ct levels across the board in my samples (higher than I would call a positive sample) and I'm wondering if this might be due to contamination when I spin down my plate. Any suggestions for spinning down plates without this issue? Would you trust a small benchtop plate spinner centrifuge to keep the wells separate?
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@Junjie Shao- No, just a sticky plate film on top.
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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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following to know
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For eDNA barcoding, is there any primer published that proved to pick seaweed rbcL genes from eDNA samples? A universal primer that could amplify rbcL from Rhodophyta, chlorophyta and Ochrophyta seaweed.
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Thanks
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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,
We are experiencing the same issue and cannot identify the problem.
We tested our primers on conventional PCR as well as the Roche SYBR green assay and both amplify with no issues. It is just with the PowerUp kit. Our primers also contain Inosine bases and we have postulated that the uracil-DNA glycosylase is the only additional component that might be the issue.
Have you been able to resolve it? Any advice would be appreciated.
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I am looking to collect various bacterial samples from plastics that have been retrieved from rivers etc. I'm looking for suggestions for the best swabs to use. I am a second year student in a cat1 lab so will not be culturing my samples. I am also looking at collecting eDNA from water and sludge etc. Any ideas for best collection methods? Thank you
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Hazel, someone in this group should be able to help / advise
John
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The main target of the survey is fish.
The location is an urban canal with a bay downstream.
I am worried about the effects of the tide.
I think whether the rising tide is good.
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Thank you for the advice.
Currently, I am reading your recommended paper.
First of all, my research will attempt both rising and falling tides.
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I am trying to figure out how many sequences I need for my samples. So far I can see that 40,000 would be necessary for microbial soil analysis. I have iDNA samples and so think I will need ~20,000 sequences however I cannot find any literature to support this.
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Hi Buffy
it depends mainly on some parameters as:
- the size of the sequence you target
- the question you want to fix (higher coverage for rare events to be detected)
- the power of the machine you'll sequence on
- the number of samples you'll multiplex
the formula can be set as:
number of requested sequence = Nb(samples) * size(targets) * coverage
fred
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Hi all,
Our lab wants to do shotgun metagenomic sequencing for environmental water samples, with novaseq 6000 system and truseq DNA PCR-Free kit (350) to make the library.
Now we have a problem that a lot of environmental DNA samples have a weak main band and strong smear, a tapesation result example is as follows.
It looks bad but we don't have enough experience. I wonder could we use samples like this to do Novaseq sequencing?
Thank you for your time.
Best,
Lingjie
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Jon Ashley Sorry for late reply. There is no specific preference for size in this project, our research is managed to look for the information from viruses to picoeukaryotes, so the target genomic range is not narrow.
We already decided to change the kit for a better preservation of sample.
Thank you a lot for reply. and wish you a happy 2020.
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How do you do to design an assay with primers and probes that function well and what program/platform do you use?
I'm using AlleleID now but I wanna try any other ways to design it.
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How can I optimeze a protocol like this? Shen-An Hwang
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When working with eDNA we sometimes face some contamination, and it may lead to false-positive or negative, and I think if some of these cases, it creates a problem, or it may be interpreted as a grown on robustness, cause if we are using universal approach, we should be capable of finding any DNA sequences, like human DNA contamination, so what do you think about it?
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Abhijeet Singh I have some other questions on my page and a discussion where there is something that I think your opinion could really help me.
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I will try to extract eDNA from water filtrated through 0.2µm membrane filter. Could I do the sequencing of 16s rRNA gene of all archaea and bacteria contained in water sample after DNA extraction?
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Yes you could sequence the eDNA using universal 16S primers.
You could also stick the filter paper on a culture plate and have a rough idea of what species are dominant in the sample, this of course leads you to isolation and study of single cultured organisms instead of the global view generated by metagenomic sequencing of eDNA from a mixed sample
good luck :)
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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!
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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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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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In this paper, they find that a comparatively simple Isopropanol DNA extraction method outperforms the kits both in terms of yield and quality. Problem is that they used intestinal tissue in addition to faeces, so results may be different if killing the fish is not an option:
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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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Environmental DNA extracted from environment samples (water, soil, gut content, etc....) while extraction DNA from specific tissue you follow the conventional method as you said from liver or flesh and so on. Now if you are using kits, you follow the manufacturer manual you can get genomic DNA. If you used the manual procedure you should check solutions conc. and pH of the analytical solution. Also, the working solution of electrophoresis pH should check
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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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You can use PICRUSt (http://picrust.github.io/picrust/), BugBase (https://bugbase.cs.umn.edu/index.html) or Faprotax (http://www.zoology.ubc.ca/louca/FAPROTAX/lib/php/index.php?section=Home). You have to use the OTU table generated with your 16S sRNA sequencing data.
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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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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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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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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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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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Thank you Waseem and Carin
Both answers are invaluable.
I remain doubtful that the barrier tips and reagents are to blame. The exact same tips and reagents were used for both treatments. The only significant difference was exposure to the air. 0 Contamination events when wells were sealed immediately with strip caps, 5 when left exposed. However, I remain open to alternatives to air-borne contamination, despite feeling the opening of post-pcr plates for purposes of capillary electrophoresis (we do not use SYBR Green to analyse melt curves) is causing the aerosolisation of DNA particles to spread throughout the lab.
I value your input regarding uv lightning, and the fume hood and laminar flow hood (the latter of which I believe has just been fitted, together with UV lightning). Indeed, when running an experiment in the fume hood, the rate of contamination was an order of magnitude less.
And yes, I have argued for the purchase of a closed-system tissue lyser from the beginning.
This contamination is quite tricky. Now looking to find statistical methods of piercing the background noise.
Thanks again
Nick
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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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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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Thanks again for the input. I'm hoping that eDNA and microbiome (bacterial) DNA will be degraded with whatever sterilization process I use so that each time I sample with the sterilized sampling equipment, there will be no worry about mixing DNA from previous collections with the new collections.
Thanks for your help.
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Did microbial DNA of sessile bacteria can integrates the genome of tumorale cells ??
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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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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 lost a lot of eDNA during this step. In addition, it cannot be ligated with the cosmid vector.
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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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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You are welcome. I hope you'll have interesting findings to share after your work. Good luck!
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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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Thank you Francesc for your advice. I will go ahead with propidium iodide then.
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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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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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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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Thank you for your answers.
The annealing recommended time is 30 seconds and extension is 45 seconds, with final extension of 5 minutes.
I do have another set of primers for that gene cluster, but the one I'm using right now is quite universal therefore I preferred to use it.
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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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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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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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you can use a temperature gradient PCR for adjusting the annealing temperature that will act with the primer and adjust denaturation at 94 degrees for 5 minutes and extension at 72 degrees for 3 minutes. the master mix will be adjusted either at 25 ul or 50 ul according to the type of the mix used and its protocol.
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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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Please see link below. It depends what your target is and the nature of it. Hope it helps:
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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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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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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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If you go for Spectrophotometer reading even for water adding some junk it gives reading.
Just load by taking 1 microliter of your DNA on the gel then do visual quantification, by comparing the readings that you got by Spec. I say that is better. Then proceed for other things like dilutions etc. etc. 
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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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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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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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A few suggestions- 
Dry your pellet in a speed vac and was it a few times with 70% EtOh, then resuspend in water.
Alternatively, a little Bovine Serum Albumin added to the mastermix can work well (it's also a bit imprecise).
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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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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 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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Thanks a lot to all those who gave me advice! I now have shortened the list of potential primers to a couple of candidates. I tried to use the package "primerTree" on R in order to compare their taxon coverage and resolution. But I didn't manage to make it work yet (got the rather obscure message "Error in `[.data.frame`(hits, location_columns) :
undefined columns selected"). Do you have any experience with this package, or with any alternative that would allow to compare primers' efficiencies for metabarcoding?
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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 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 Jon,
I think you are mentioning about recovery of the bound aptamers from the membrane?
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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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Dear Paul,
I'm not sure I understand your question. Are you looking for a database that can match your EDNA sequences to meiofauna species, or just a list of species that belong to meiofauna?
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