Science topic

Molecular Ecology - Science topic

Molecular ecology is a field of evolutionary biology that is concerned with applying molecular population genetics, molecular phylogenetics, and more recently genomics to traditional ecological questions (e.g., species diagnosis, conservation and assessment of biodiversity, species-area relationships, and many questions in behavioral ecology).
Questions related to Molecular Ecology
  • asked a question related to Molecular Ecology
Question
4 answers
To better grasp fundamental evolutionary concepts and models, as well as the past and most recent techniques of sequencing. Ideally freshwater & marine ecosystems are one of the focus.
Thank you for your recommendation!
Relevant answer
Answer
Molecular Ecology, 3rd Edition
Joanna R. Freeland
ISBN: 978-1-119-42615-8 January 2020 384 Pages
  • asked a question related to Molecular Ecology
Question
8 answers
I'm trying to carry out the Mantel test using Arlequin, has anyone here done it before?
Relevant answer
Answer
For the mantel test, you need to use the arp file attached here
Thanks
  • asked a question related to Molecular Ecology
Question
3 answers
So, we started using silicone sealing mats on our plates for the PCR diagnostics. They seem to fit well and there is no leakage. The problem is, they often tend to pop out during the PCR run, causing all the water to evaporate. We figured that this happens very early, during the initial activation. Any idea why this might happen? Thank you
Relevant answer
Answer
Hi, Lara!
Maybe I can help.
If I understood well, you are using a 96 well plate and you place this well into a PCR machine? And after that, you seal it with a silicone mat? And your PCR protocol has a hot-start period where you need to activate the polymerase enzyme?
If everything mentioned above is the case, then I am afraid that using these silicone mats could, unfortunately, be the problem. Especially during the initial activation, where you are holding the samples at a high temperature for a longer time and the pressure is building up inside each well, resulting in the silicone mat popping out.
Additionally, this can be an issue due to the 96 PCR well being too smooth as well. But I wouldn't worry about the PCR plate as it is important that it is smooth so that DNA doesn't stay on it.
There are several ways how to avoid this:
1) Try to buy some other sealing mats to see if they work better with your 96-well plate. Or try buying new mats and plates together from the same company if you already didn't do that.
2) Use a PCR kit and protocol without a hot start, although I wouldn't recommend that as it could potentially affect your primer specificity and lead to primer dimers.
3) Try to buy standard 0,2 mL PCR strips for 8 samples in a row. Something like this:
I use such strips for my PCR as I have more control with opening and closing these tubes and using them at once only as much as I need. These strips never disappointed me and never popped out while performing PCR, and I did it with them in 5 different PCR machines that our lab has to offer.
I hope this was of some help.
Best regards,
Domagoj
  • asked a question related to Molecular Ecology
Question
6 answers
I have a dataset of 12 nuclear markers. Can I conduct population analysis (AMOVA, Fst) using Arlequin by multilocus sequence nuclear data? How can I prepare the input file?
Relevant answer
Answer
Hi Larissa
I'm having the same problem as you had, with 5 loci.
I tried to change the Data type: Standard, and use the haplotype data, not the sequence, but I'm not sure if it works the same with the sequence.
  • asked a question related to Molecular Ecology
Question
4 answers
I've tried several programs (e.g. TreeView, FigTree, MEGA, Archaeopteryx, Mesquite) but they just display node numbers and branch lengths. I'm using the software SYMMETREE to infer diversification rate shifts on particular nodes within a tree. Results refer to "branch numbers" which, according to the manual, can be displayed using MacClade. However, I´m not using a Mac OS platform.
Relevant answer
Answer
Yes, ape in R is a good option
For the sake of providing a direct answer to the SPECIFIC question you asked, the following R code will do exactly what you require, and output the tree to a PDFfile in your current working directory
library(ape)
tree = read.nexus(file="path\to\file")
tree_export = "tree.pdf"
pdf(file=tree_export, width=6, height=6)
plot(tree, cex =0.5, use.edge.length=FALSE)
axisPhylo()
edgelabels(cex = 0.25, width = 0.1)
dev.off()
depending on your tree structure just play around with the "cex"and "width" parameters to make the branch numbers readable, the above peramters are reasonable starting points.
The above code assumes your tree is in Nexus format. For the commonly used Newick format do this:
library(ape)
tree = read.tree(file="path\to\file")
tree_export = "tree.pdf"
pdf(file=tree_export, width=6, height=6)
plot(tree, cex =0.5, use.edge.length=FALSE)
axisPhylo()
edgelabels(cex = 0.25, width = 0.1)
dev.off()
i.e. swap "read.nexus" for "read.tree" function
  • asked a question related to Molecular Ecology
Question
3 answers
I would like to find a tool (if it exist ...), to predict the porportion of r vs K strategist in samples from a metabarcoding study.
For example, the tool I need (R package or equivalent) could work similarly to functional predictive tools such as Tax4Fun, FAPROTAX or PICRUST, but instead of predicting functions, it investigate ecological strategy such as r vs K as explained by the "r and K selection theory".
Benoît.
Relevant answer
Answer
Hi Benoît,
No program that I am aware of. I reckon that differentiate life-strategist from mere amplicon seqs can be, at best, unreliable if considering problems such as uncomplete reference datasets.
A good paper describing this but trying to list taxa that behaves as copio-oligotrophs:
Please let me know if you come across with a relevant program though
Cheers
Ben
  • asked a question related to Molecular Ecology
Question
6 answers
Hello People
I am extracting host DNA from fecal samples. However, feces contains lot of microbes and their DNA too gets extracted during DNA extraction procedure.
I want to get rid/minimise the microbial DNA. It is causing a lot of issues in my PCRs (lots of misamplification). Is there an inexpensive method to do so?
Relevant answer
Answer
Yes this question can really only be addressed in a helpful manner with more information about your needs and unfortunately your budget. But basically eDNA techniques are emerging that allow simultaneous amplification/sequencing of all of the OTUs that successfully amplify, so by definition this approach uses universal primers for markers usually such as 16S rRNA and ITS, internally transcribed spacer region, or COI cytochrome c oxidase I, then the resulting sequences are blasted against NCBI database to determine biodiversity in the sample. But if you are using a more traditional PCR approach where you want to specifically amplify only certain target lineages, one possibility is use species of specific primers. Depending on the species of interest, this can be tricky if such PCR primers are not available, but you can design your own primers that specifically amplify only the species of interest, provided there are DNA sequences available for these taxa.
  • asked a question related to Molecular Ecology
Question
2 answers
Dear colleagues,
Unfortunately, the “Permanent Genetic Resources” of the journal “Molecular Ecology Resources” are not permanent, because the database (http:// tomato.biol.trinity.edu) no longer exists. The journal editor could not help with that issue, it seems that all the “Permanent Genetic Resources” published in papers of the journal “Molecular Ecology Resources” that have been deposited in this database are lost. This raises questions about the current trend of relying solely on online publications, but that’s a different story.
I am looking for information on the described microsatellites and primers of the species Thais(ella) chocolata from the following publication:
Permanent Genetic Resources added to Molecular Ecology Resources Database 1 December 2010–31 January 2011. Molecular Ecology Resources (2011) 11, 586–589
The microsatellite sequence is published in GenBank, but not the primers.
We have contacted already one of the authors, but we didn’t receive the primer sequences.
My question is, if perhaps someone had retrieved detailed information about the microsats and primers from Thais(ella) chocolata before the database was shut down.
Thank you very much in advance.
Best regards,
Marc
Relevant answer
Answer
How much of the sequence is published? If its just the microsatellite region, then you might be out of luck. If there is sequence flanking, you can design your own.
  • asked a question related to Molecular Ecology
Question
19 answers
I am enjoying expanding my molecular tool belt by included SNP markers on a non-model plant species. What I would really like to find out from this group of outstanding scientists is a data analysis package that can handle SNP data. Yes I am aware of a work around for Arlequin, but I would prefer to actually use a program written for SNP data. Suggestions would be helpful.
GenAlEx, Structure, and PolyR have pros and cons (see my Aug 28, 17 answer/update) Danny
Relevant answer
Answer
This website might give you some initial lights:
Also, note it has a arXiv paper associated to it.
  • asked a question related to Molecular Ecology
Question
13 answers
I have a dataset of 6 microsats loci from 11 populations of a barnacle. I’d need 100-1000 matrices of pairwise Fst-values, in order to carry out a spatial analysis. Is there any software producing these bootstrapped matrices?
Relevant answer
Answer
I had some troubles running diveRsity::writeBoot.
diveRsity::basicStats and diveRsity::diffCalc ran my data. However, I had to make minor adjustments to my data to writeBoot (remove some empty spaces).
Also, parallel argument must be set to TRUE. parallel = FALSE causes problems.
  • asked a question related to Molecular Ecology
Question
6 answers
I'm trying to run a scenario with two populations. I think it might have got something to do with the conditions set on the time and Ne parameters, but I'm stuck. Thanks in advance!
Scenario:
N1 N2
0 sample 1
0 sample 2
t1-db varNe 1 N1b
t2 varNe 2 N2
t3-db varNe 2 N2b
t4 merge 1 2
Conditions:
N2>N1b, N2>N1, N1b<=N1, N2b>N2, t4>t2, t4>t3, t4>t1, t3>t2, t2>t1
Error:
Something happened during the reftable generation :
Program of thread 'guanacos reference table generation' exited (with return code 1) unsuccessfully.
I will try to save current RNGs' states.
I have saved current RNGs' states into /Users/ana_agapito/Documents/Documentos/ubb/paper MSS/lamaguanicoe2/guanacos_2018_8_3-1/RNG_state_0000.bin
Content of progress file :
Relevant answer
Answer
Hi Ana,
In the scenario you specify "t2 varNe 2 N2" but "N2" is already the effective population size specified on line 1. So this line is not really changing the Ne of population 2 like you might want it to. Try changing it's name to see if this solves your problem. Also make sure the naming of all variables is consistent. You have t1-db in the scenario but just t1 in the conditions.
Also, for your conditions you can also remove some redundancies. For example, t3>t2 and t4>t3 imply that t4>t2 and so you could remove this condition. For the same reasons N2>N1b and t4>t1 are redundant conditions and could be removed as well.
Hope this helps!
  • asked a question related to Molecular Ecology
Question
3 answers
I am interested how fast can the cluster membership change. I know it depends on the parental genotypes, but how strong this change could after one or more generations? Im working with Q-values from Structure software on Rousettus aegyptiacus population structure. It would be helpful if anybody could recommend some literature.
Thanks!
Relevant answer
Answer
A paper worth reading (linked below) illustrates that the time lag between an isolation barrier and detecting this pattern via genetic clustering methods (eg. STRUCTURE or TESS) can be as low as 15 generations for organisms with relatively large dispersal abilities. In other words, impacts to genetic clusters can be altered relatively quickly, which is especially pertinent to your study organism. Hope this helps.
  • asked a question related to Molecular Ecology
Question
20 answers
I am currently studying hybridization between 2 plant species using dominant genetic markers (ISSR markers), and would like to infer the stages of hybridization (infer with confidence that a certain individual is an F1 hybrid, F2, or backcross). So far the only one I have found is NewHybrids by Anderson and Thompson 2002 (http://ib.berkeley.edu/labs/slatkin/eriq/software/software.htm). Does anyone know if there are other softwares that could do a similar job, probably with a different model or inference method?
Relevant answer
Answer
For those with access to a Mac with OSX, the following may be helpful, as it appears the Mac version is updated more often. I was able to get NewHybrids (without graphics) to run reliably using the steps below. Note that I'm no programming expert, so there may be easier ways, but this worked well for me after trudging through available documentation online. Hopefully this saves someone some hours of agony.
1) Download and extract all files from GitHub site: github.com/eriqande/newhybrids
2) Follow folders system to Bin -> OSX -> newhybridsng.exec (run this; no graphics version)
3) Back at the main folder downloaded from GitHub, open a new Terminal Window at this folder location (can be done on some machines by right-clicking the main folder)
4) Copy and past the following into the terminal to see a helpfile of all available commands: ./bin/OSX/newhybsng --help-full
5) Using this documentation, string together necessary analysis of desired datafile. Note that "--no-gui" command is needed at end to make the "no graphics" version work. The below example loads the test data file "TestDat.txt" that comes with NewHybrids, along with personalized model specifications. Also note that your data files are likely stored in a different location, but you may be able to drag-and-drop the data file into your Terminal window to obtain the file path needed.
Example Terminal command: ./bin/OSX/newhybsng --datafile /Users/Andrew/Desktop/newhybrids-master/test_data/TestDat.txt --theta-prior uniform --pi-prior uniform --burn-in 50000 --num-sweeps 100000 --no-gui
  • asked a question related to Molecular Ecology
Question
3 answers
Is there an application of comparative parallel application of methods used in Ecological PopGen (Ie invasive species).
Can such measures of gene flow, population expansion, AMOVA's, Tajima's D, raggedness etc be applied to Cancer or other diseases. Moving from Ecology into BioMedical Science?
Relevant answer
Answer
A combination of haplotype grouping, network analysis of genes interaction and population genetic background of affected paitients could produce interesting results. We have initiated this type of study with male infertlity in our country, I hope in comming 2 years we can get some introductory results and insights to this hypothesis.
  • asked a question related to Molecular Ecology
Question
10 answers
Dear colleagues,
I am interested to see how individual genetic diversity changes in the landscape. My individuals are spread in a large landscape and it doesn't make sense to assign them to populations. There are some ways to calculate individual genetic diversity (see Aparicio et al. in Molecular Ecology (2006) 15, 4659– 4665). Here comes the next problem: my species is tetraploid. Does anyone have an idea?
Thanks in advance,
Charalambos
Relevant answer
Answer
  • asked a question related to Molecular Ecology
Question
4 answers
Here is my question:
I have used several kinds of genetic markers to reconstruct the phylogeographic history of a kind of bats. A geographic relatively isolated population (call it YM population) contained three adjacent phylogeographic clades inference from mtDNA with lowest nucleotide diversity and haplotype diversity in both mtDNA and ncDNA, and firstly isolated from other populations in microsatellite analysis. ncDNA suggested weak geographical pattern of all clades inferred from mtDNA.
I want to figure out the potential origin of YM population, or the possible reasons for low nucleotide diversity and haplotype diversity in both mtDNA and ncDNA. By the way no bottleneck (based on microsatellite) or expansion (based on mtDNA) was detected.
Any suggestion will be appreciated including the question itself.
Relevant answer
Answer
Depends on the number of loci and quality of the data. IMA2 and DiYABC requires many loci and high quality of the data. Use different bottleneck tests. To check for contemporary bottlneck you can try Wang's Colony. 
  • asked a question related to Molecular Ecology
Question
19 answers
While calculating the evolutionary divergence as computing pairwise distance, (using p-distance methods and Maximum Composite Likelihood model), the values ranged between 0.002-0.138 (p-distance) and 0.001-0.109 (ML) for between individuals of the same species, and for different species. I have got these value as distance matrix table using MEGA 7 software by inserting the nucleotide sequence of COI gene of 661 bp length. Can somebody tell me what are the actual threshold values of the evolutionary divergence calculated by the above said methods to delimit the distinct species? I mean at which values of these distances we would say that these two species/individuals are of distinct species?
Relevant answer
Answer
I would emphasize even more strongly than the previous respondents that speciation is a process where subpopulations establish their own independent evolutionary trajectories, and any notion of a threshold difference is only a rough approximation at best.  As an example, back in the '90s I was doing a postdoc where I was sequencing mtDNA of macaque monkeys.  At that time we discovered that rhesus monkeys had major haplotypes differing by about 3%.  A very well known evolutionary biologist from the American Museum of Natural History assured me that we had made a mistake because the threshold for species elimination was 1% for mtDNA.  Since then a different species of macaque has been found to contain mtDNA sequences that differ by about 9%.  Macaques have relatively extreme female philopatry, which explains this unusual observation, but it highlights the folly of assuming an arbitrary threshold of difference.
  • asked a question related to Molecular Ecology
Question
7 answers
Does anybody have idea to determine polymorphic bands in a binary matrix (ISSR) and how to calculate the genotypes provided from each locus. I calculated the number of bands which present 0 and 1 in the different isolates and I divided it by the total of bands (I considered the bands that gave 1 for all isolates as monomorphic band) I found 80 % of polymorphism however when I constructed the dendrogramme (BY UPGMA) it show a very similar structure for the majority of isolates with low variability and low valor of boostrap. I can't see the logic of this result. ..Thank you for your help. 
Relevant answer
Answer
Hi, 
For dominant markers like ISSR, you can determine many genetic diversity parameters like Nei gene diversity, Shanon information index, effective No. of alleles, etc. Please read the paper attached and other papers published by our team. 
You van use Genalex and Popgun for such analyses. 
Regards 
Masoud Sheidai 
  • asked a question related to Molecular Ecology
Question
8 answers
Hi everyone,
I would be interested in your opinion about most widely used metrics to assess genetic diversity, particularly when considering markers specific features.
Expected heterozygosity, allelic richness and privates alleles are common in microsatellites-based studies for example. It seems He/Ho and nucleotide diversity are widely used for high-density SNP data, haplotype diversity providing additional information if GBS is used.
However when working with medium density dataset (~100 SNP) without any knowledge about the genome and no sequence data, I feel like classical measures like He or polymorphism rates might be tricky to compare, for example if I want to compare genetic diversity between two geographic regions. I am afraid to be lacking of statistical power, especially when number of individuals in a location is also low;
Would you have any idea/personally recommand a particular approach to this end ? Could you recommand any paper about study of genetic diversity of SNP aside from HTS ?
I thank you very much for your answers and wish you a very nice day :)
Chrys
Relevant answer
Answer
using 100 SNPs (assuming a reasonable minor allele frequency) is not such a difference in resolution compared to classical AFLP studies. 
gene diversity is nothing else than the expected heterozygosity (He). If you have an average gene diversity of c. 0.15 - 0.20 (which is the chance that if you take two random alleles from your sample they are different), you're in the typical range for this type of population genetic studies. 
In the early days, many allozyme studies had similar diversity ranges (but fewer loci). So it's pretty OK to use He, compare it to Ho, calculate allelic richness, and so on. The more loci, the smaller the error on the estimate of genome-wide diversity and so on. So more loci allow you to discern small effects from noise better. 
Basically, all classical population genetics is based on detecting deviations from Hardy-Weinberg expectations (F-statistics at different hierarchical levels) to test for evolution. (the HW theorem assumes no evolution, hence it is used as a null model). Any deviation from HW is caused by evolution, and the trick is to disentangle the three (four) major drivers of evolution: drift, gene flow (and subdivision), selection (which is different per locus due to recombination) and mutation (which is in itself typically negligible compared to the three other).
More importantly, what questions are you trying to answer? Do you require individual-based analyses (assignments, clustering, parentage, ...), are you satisfied with estimates at the population level (summary statistics), etc.
As a reviewer, I'd rather read a manuscript with a good sampling design for the question asked, crisp statistical analyses, clear knowledge on the assumptions of each analysis (the biggest issue in population genetics if you ask me), and using only 100 SNPs, than a study that uses 100 000 SNPs but has no clue how to really answer the question (other than "we were interested in ..."), other than by blindly copying what others have done in somewhat similar studies.  
  • asked a question related to Molecular Ecology
Question
3 answers
Hi all,
I have pooled RAD-seq data (multiple individuals sequenced in a single library prep) and I am interested in haplotype frequencies.
Reads have been mapped to a de novo reference and I have a BAM file for each population.
It seems reasonable to assume that haplotypes frequencies could be called from BAM files in a manner analogous to determining allele frequencies in sequenced pools; e.g. counts of different haplotypes in the sample.
However, I cannot seem to find any good software specifically designed to deal with pooled RAD data. GATK and HaploPool, for example, are extremely limited by ploidy size. Collen Beck's rad_haplotyper (https://github.com/chollenbeck/rad_haplotyper) is nice to use and good for RAD data but assumes individuals as the sequenced unit; at most it would only provide information about which populations are likely fixed for particular haplotypes.
Does anyone have good recommendations?
Cheers,
~ Josh
Relevant answer
Answer
@Ivan and @Yann: Thanks for comments. I am using Freebayes to call SNPs because it has an argument that considers the mapped BAM files as deriving from pooled samples. BUT I REALLY WANT HAPLOTYPES!
Also, I have worked with the Popoolation tools before, but my understanding is that they are better suited to projects where you have much larger contigs (e.g. choromosomes as opposed to RAD fragments).
It confuses me that so little has been done to achieve this gap in analysis tools.
  • asked a question related to Molecular Ecology
Question
5 answers
After the following two articles, I got confused and am trying to figure out whether the method of DNA barcoding gap to identify species is suitable or not. How should i treat this method?
Srivathsan, A., & Meier, R. (2012). On the inappropriate use of Kimura‐2‐parameter (K2P) divergences in the DNA‐barcoding literature. Cladistics, 28(2), 190-194. DOI: 10.1111/j.1096-0031.2011.00370.x
Klemen Candek and Matjaz Kuntner. (2015). DNA barcoding gap: reliable species identification over morphological and geographical scales. Molecular Ecology Resources (2015) 15, 268–277. DOI: 10.1111/1755-0998.12304
Relevant answer
Answer
Barcode Gap is a suitable method to delineate species from a set of aligned sequences as long as you have enough sequences per species. Yet, it is important to remind that Barcode Gap should not be the only criterion! Complementary evidences like other genetic markers, morphology and distribution are necessary to make a decision. I recommend reading the ABGD method, see link attached.
Once you have established a reference dataset with barcode gap then you can be confident with the identification of an unknown specimen through it's barcode sequence.
Hope that helps.
  • asked a question related to Molecular Ecology
Question
6 answers
I am investigating hybrids between two plant species. The parent of one has a strong sulphurous smell, and this is passed onto the hybrids. 
We are looking at the morphological and genetic differences, but it would be great to also see the chemical differences.
Can an HPLC analysis be used to quantify the sulphurous compound across samples? (We have an HPLC machine in the department, but I have never used and this would be a new use for it so the technicians are unsure). If yes, any suggestions or pointing to relevant literature or methods descriptions would be greatly appreciated.
Thanks in advance for any suggestions.
Kind regards,
Alastair
Relevant answer
Answer
Thanks Bruce for the very helpful answer about Mass Sectrometry.
Kind regards
Timothy
  • asked a question related to Molecular Ecology
Question
2 answers
Hi all,
Thank you for your help with this.
I'm using Bayenve2 to identify loci under selection across a climate gradient
What you get with bayenve2 is quite a large number of false positives, where the high BF loci can be due to reasons such as a single population having an extreme value and genotype, thus the GxE correlation is created by a single population rather than correlation over a consistent climatic gradient.
It seems potentially that one needs to visualise the data for each putative loci under selection to review whether it is due to a single population or the required allele fluctuations over a climate gradient.
Given that Bayenve2  internally calculates the degree of allelic frequency due to environmental cause, over demographic history, is there anyway to extract and just visually plot the  corrected allele frequencies due to environmental affects cross populations to show the potential linear correlation between allele fluctuations and for example temperature?
Cheers
Jamie
Relevant answer
Answer
Hey James,
Not sure what you mean by 'corrected allele frequencies' but with BayeScEnv2 you can actually identify the putative loci under selection (I think it calculates local Fst and correlates this measurement of genetic differentiation with the environment). So you can always extract a data set with only the loci under selection and plot them. Is this answering your question?
  • asked a question related to Molecular Ecology
Question
3 answers
Currently, I'm trying to make plant phylogeny reconstruction using Bayesian inference and have a need in applying two different evolutionary models for different parts of sequences in one sampling because they have a secondary structure with paired and unpaired regions (ssRNA and dsRNA) which has an influence on nucleotide substitution frequency. I want to take into account such impacts. Is it possible to calculate Bayesian inference for phylogenetic purposes using two different evolutionary models with different parameters for different parts of one sampling of sequences simultaneously?
Thank you in advance.
Relevant answer
Answer
Yes, you can divide your dataset into partitions and apply different sequence evolution models to those partitions. A common framework might first involve using PartitionFinder (http://www.robertlanfear.com/partitionfinder/) to delineate partitions and their respective "best" models.
Sounds like you are using Mr. Bayes? You would then define your partitions in the Mr. Bayes command block of your nexus file.
For some help in configuring your nexus file for a partitioned dataset, see: http://mrbayes.sourceforge.net/wiki/index.php/Analyzing_a_Partitioned_Data_Set
  • asked a question related to Molecular Ecology
Question
4 answers
I wonder whether substitution saturation can influence only the results of phylogeny reconstruction, or whether it can also influence results of DNA-based species delimitation and/or DNA barcoding. Does anyone know? Or what is your opinion? I will appreciate any contribution on the topic.
Relevant answer
Answer
Biologically, the probability of mutation occurred at the same site more than once (multiple hits) is small (given the low mutation rate), the lower mutation rates the smaller the likelihood given a time interval. Given a certain mutation rate, the smaller the time interval the less likely of substitution saturation.
  • asked a question related to Molecular Ecology
Question
4 answers
Hello,
I would like to try the trypan blue to stain dead organisms. However, I tried to prepare the stock solution of trypan blue at 0.4% with a powder of trypan blue, and PBS, and even if I filtrated the solution, it precipitated. I don't understand why. Does someone have a good protocol to prepare a solution of trypan blue, and who do you use it to stain dead organisms?
Relevant answer
Answer
Hi 
you can use this protocol:
Making a stock solution of 0.4% Trypan Blue (also available for purchase):
Weigh 4 mg of Trypan Blue powder and add 1 mL of water or 1x PBS. Scale up as necessary
 
Using Trypan Blue to stain dead cells in tissue sections:
Mix 1:1 the Trypan Blue solution with 1x PBS
Add 1mL of this dilution to tissue sections, mix well, and incubate at room temperature for 5 minutes
Rinse tissues with 1x PBS for 15 minutes
 
Using Trypan Blue to stain dead cells in suspended cultures:
Dilute your cell suspension in a 1:1 dilution with the 0.4% Trypan Blue stock solution
Carefully fill the hemocytometer chamber with the stained cells
Incubate the cells for 1-2 minutes at room temperature and proceed with counting/quantitation
  • asked a question related to Molecular Ecology
Question
1 answer
I am having trouble finding non-coding cpDNA regions that will provide enough variation and clean reads for for phylogentic analysis. As a result I only have three D4 introns to use. Usually I would have some intergenic spacers or introns from other areas of the chloroplast genome as well.
Is it viable to combine and analyse sequences from the D4 set alone? Or does that result in a high risk for some sort of evolutionary linkage due to their close proximity to one another?
Relevant answer
Answer
Hi,
See the link below. 
Good luck. 
  • asked a question related to Molecular Ecology
Question
10 answers
Trying to get Powerstats version 1.2 (Promega Corp.) software, but is is not available. how can I get the software to calculate power of discrimination (PD) and a prior exclusion chance? The STR population genetic statistics can not be calculated with other way.
Relevant answer
Answer
Hi Shiva,
Did you find the software? If not how did you calculate PD or PE?
  • asked a question related to Molecular Ecology
Question
10 answers
I am trying to see whether my data from 45 stands of pedunculate oak (Σ=1120 individuals) are structured. Using STRUCTURE and BAPS, I could not get any strong evidence about genetic structure (maybe only some vague hints; see previous question).
After recommendations of you, I carried out a DPCA (discriminant analysis of principal components using the adegenet package in R). The BIC vs. K diagram suggests K=7, although it's suspicious how small differences in BIC among different values of K are. The scatterplot looks nice and suggests subdivision, I think, although overlapping is also present. On the other hand, a bar plot of membership proportion does not reveal any recognizable pattern. All seven clusters are more or less equally represented in each population.
Here is a link with diagrams (membership proportion is shown only in one example population, but others look similarly): https://drive.google.com/folderview?id=0B41pt4LUC4K3S2xnUXFsaWRHanM&usp=sharing
I'm not experienced in this method. Can anyone have a look and tell me her/his opinion whether (1) the method indeed suggests genetic structure, (2) whether also other K values should be used.
By the way, I followed the instructions from the manual to decide about the number of principal components used: 500 for "find.clusters" and 200 for "dapc", "to avoid over fitting".
Thanks in advance for your help.
Relevant answer
Answer
Actually the structuring phase comes after to check the IBD (isolation by distance). For IBD the most commonly use criteria is Mental test but their is also some biases (Please read: Meirmans PG (2012) The trouble with isolation by distance. Mol Ecol 21:2839–284), so I would like to suggest first do IBD through RDA (Meirmans PG (2015) Seven common mistakes in population genetics and how to avoid them. Mol Ecol 24:3223–3231), if their is no IBD then go through the clustering, may be in your case you have  just one genetic group in actual not more than one.
  • asked a question related to Molecular Ecology
Question
3 answers
How can I consider together the low genetic diversity (haplotype diversity:0.0000 to 0.042 and nucleotide diversity: 0.00000 to 0.00089) and high FSH values (-0.0546 to 0.29807) seen in marine fish species.
Relevant answer
Answer
Sorry,
I wanted to write Fst (pairwise)
Best regards
  • asked a question related to Molecular Ecology
Question
8 answers
Hey lab folks, for PCR do you prefer disposable (plastic or metal, one time use like https://www.fishersci.com/shop/products/corning-sealing-mat-384-well-polypropylene-storage-plates/07202500) or reusable (usually silicone, washable and autoclavable like https://www.fishersci.com/shop/products/axygen-impermamat-axymat-sealing-mat-2ml-96-well-deep-well-plates/14222010) type plate sealers? Which would reduce contamination the most?  I will be doing microsatellites for the most part. 
Relevant answer
Answer
I use reusable plate sealing mat for my PCR. I have not faced contamination problem as far, even after reusing the mat multiple times. 
I would recommend reusable plate sealer but care should be taken while washing and autoclaving to avoid contamination. 
  • asked a question related to Molecular Ecology
Question
3 answers
I have used STRUCTURE and BIMR to describe population structure and migration rates. But, now I want to identify individual migrants within these sampled populations.
Relevant answer
Answer
yes there is a "posteriori" assignment test in Geneclass2. I've run it LOOOONG time ago so I'm not sure how it is in the current version, but it wasn't hard to interpret at all. The main question is to have "GOOD reference populations" to assign these potential immigrants to. So what I was suggested to do when I've applied it, is to use as reference pops the genetic clusters resulting from STRUCTURE. Thus, the likely immigrants would be like "ouliers" if you can say that way from those in theory "panmitic" genetic units resulting from Structure. But is fast to run, so you can run both with the genetic clusters and the populations they were captured in....  and interpret the results  from both assignments.
  • asked a question related to Molecular Ecology
Question
4 answers
Warm Greetings, Fellow Researchers!
I am currently doing research on ocean acidification and its effects on corals in the Philippines. I would like to determine coral colonies resilient to ocean acidification for the purpose of developing appropriate and efficient restoration techniques. I am currently looking into molecular markers that may serve as basis for stress response and in turn resilience, of the coral colonies. I am currently thinking of microsatellite genotyping. Would this be the best approach? Thank you!
Relevant answer
Answer
Mr. John,
Thank you for your answer. I will look into heat shock proteins right away. Ideally we might be able to study the entire genome so I will look into that as well!
Many thanks,
Allan Copuyoc
  • asked a question related to Molecular Ecology
Question
4 answers
What is the nature of inserts between the forward and reverse priming sites? What will be the properties?
Relevant answer
Answer
Hey! My question do you want to sequence the SSR product? If yes its fine, I answer your question. Why the question of evolution come here, amazing and interesting answers!! 
  • asked a question related to Molecular Ecology
Question
3 answers
If it is , then can anyone mention an example in plants?
Relevant answer
Answer
What do you mean by linkage inheritance? Genes are linked and are inherited together both in plants and animals.
  • asked a question related to Molecular Ecology
Question
5 answers
Even though we had an expected product size in SSR amplification sometimes more number of bands are visible in agarose gel electrophoresis;How can we avoid this?
Relevant answer
Answer
If you are sure for experimental conditions suggested by Dr. Pandit and Dr.  Ramadevi, It means that your primers targeted more than one loci due to paralog loci . In such a case you can score PCR products desperately for mapping such as SSR1a and SSR1b.  
  • asked a question related to Molecular Ecology
Question
8 answers
I am working on molecular phylogeny in the order level. When I am extracting the sequences from NCBI Genebank, I found there are lot sequences marked as unverified. For concratenation of multiple genes, there are some sequences are missing or unverified in some species. May i take those for my phylogenetic analysis? Are there any problems in those sequences?
If there is no problem, then why those unverified sequences are present in NCBI Genebank??
Relevant answer
Answer
 I am quoting gene bank
"As part of the standard submission process, GenBank staff review submissions for biological accuracy and assist authors in providing accurate annotations. If GenBank staff is unable to verify the accuracy of the submitted sequences and/or annotations, they may now add a comment to the record stating that the sequence is unverified. Until the submitter is able to resolve the issues, such sequences will have the word ‘UNVERIFIED:’ at the beginning of their definition lines and will not be included in BLAST databases."
Shefali
  • asked a question related to Molecular Ecology
Question
5 answers
I am working on conservation genetics of some alpine medicinal plants. I have used some dominant markers and co-dominant markers. I am unable to construct network tree by using Splitstree.
Can anybody please help me to construct the input matrix for "Splitstree" or "Neighbournet" using binary data matrix (1,0).
Relevant answer
Answer
Hi
You can get binary data, such as for dominant markers like AFLP, into splitstree using this format
#NEXUS
BEGIN Taxa;
 [&&SPECTRONET SN_BLOCKNAME Taxa_2]
 DIMENSIONS NTAX=4;
 TAXLABELS tax1 tax2 tax3 tax4;
END;
BEGIN Characters;
 [&&SPECTRONET SN_BLOCKNAME Characters_1]
 DIMENSIONS NCHAR=10;
 FORMAT DATATYPE=STANDARD GAP=- SYMBOLS = "012345";
 MATRIX
tax1 01010 01010
tax2 11001 00101
tax3 01110 11001
tax4 10101 10101;
END;
For co-dominant markers, you can code the data as binary data by recording the presence/absence of each allele - but this is not a particularly good approach. If you want to combine dominant and co-dominant markers together it will function.
For co-dominant markers alone, it is generally better to use a population genetic software package to generate a distance matrix for the samples/populations and then get that into Splitstree using the format you can find in the examples folder supplied with the software in the file south.nex. Be careful to use an appropriate genetic distance measure in your population genetic software! My experience would suggest being very wary of models that do not treat all alleles as equally related. Typically the Nei (1972) model is a good start.
  • asked a question related to Molecular Ecology
Question
7 answers
Which are the best PCR primers for Citochrome Oxidase I gene (COI) specie identification of mammals ? and birds?
Thanks in advance
Relevant answer
Answer
The following primers give very good results for a large array of bird species:
CO1-Ext Forward: 5’- ACGCTTTAACACTCAGCCATCTTACC-3’
FISH1Reverse: 5’–TAGACTTCTGGGTGGCCAAAGAATCA-3’
annealing temperature=54°C
Refs:
Johnsen, A., Rindal, E., Ericson, P.G.P., Zuccon, D., Kerr, K.C.R., Stoeckle, M. and Lifjeld, D. 2010. DNA barcoding of Scandinavian birds reveals divergent lineages in trans-Atlantic species. J. of Ornithol. 151: 565-578.
Ward, R. D., Hanner, R. and Hebert, P. D. N. 2009. The campaign to DNA barcode all fishes, FISH-BOL. J. Fish Biol.74: 329-356
  • asked a question related to Molecular Ecology
Question
3 answers
interstitial,karachi
Relevant answer
Answer
thanks Melih
  • asked a question related to Molecular Ecology
Question
5 answers
I am looking for rates to calibrate my analyses, but have not found any so far. Does anyone know of any research done in this area? There appears to be plenty on mtDNA markers, microsatellites and some nuclear genes, but none on neutral nuclear markers. Any help would be much appreciated!
Relevant answer
Answer
Hello Ashley,
It depends on the loci you are using and the group you are examining (hard to generalize across all fish). I have examined several markers in Centrachidae in the southeast US and Catosomidae in the southwest US and here are some insights that I have found.
Both of these groups have very differing substitution rates. Centrachidae has substitution rates similar to that of the average mammal rates, reflecting and more stable history with relatively large populations consent throughout time. Catosomidae, like most western fishes, have had an unstable history driven by volcanism and continental expansion and as such has presumable had periods of bottlenecks and expansions over and over again leading to a quicker substitution rate (almost an order of magnitude), this was found in Chen and Mayden 2012 and is consistent with the work I have in review. So history is very important and other factors including introgression, genome architecture, and genome duplication can also be very important.
Also, as you mention mutation rates vary across the genome. Even 'neutral markers' are not completely neutral but instead nearly-neutral and many of the markers that we call neutral are not. The general rule of thumb for eukaryotes (although these may be more basis to mammals) is an average of 2.2 x 10^-9. Below is a general list for various regions of the genome:
Protein-coding exons - 10^-9
Introns - 10^-8
Ribosomal proteins (like S7 that Derek mentioned) - 10^-6 to 10^-7
Microsatellites (and many other short tandem repeats) - 10^-4 to 10^-6
Mitochondria - 10^-5 to 10^-6
There are also ultra-conservative regions that have a much slow mutation rate and have become popular in the NGS world. Also, when people refer to neutral markers they often are referring to either ribosomal proteins or EPIC markers (exon primed intron coding) that often contain both exons and introns and when you sequence these markers you can often see the rate heterogeneity.
With all that said, applying a standard rate is possible and many researchers do using the standard eukaryote mutation rate of 2.2 x 10^-9 (can not think of the citation right now), but you should be careful and only use it on EPIC markers and not other markers like ribosomal proteins (i.e. S7 and 18S). Also this should be avoided if your organism has had a history of bottlenecks and expansions. The far better thing to do in my opinion is to fossil calibrate your marker as long as you have the calibration points. For examples in fishes see Near et al., 2012 in PNAS for all ray-finned fishes or Unmack et al., 2014 in PloS for Catosomus.
Hope this helps. There is definitely literature out there, they can just be hard to find and typically easier to find if you look more at the family level then in all of fishes.
Best Regards
  • asked a question related to Molecular Ecology
Question
11 answers
I am trying to get a dendrogram from microsatellite data. I am comparing among individuals, not populations. I got a dendrogram from MEGA, by using a genetic distance matriz generated in GENALEX but i am wondering if si possible to get a dendrogram with boostrap numbers.  Thanks in advance!
Relevant answer
Answer
DARwin 6.0.12
  • asked a question related to Molecular Ecology
Question
7 answers
I found this interesting paper that reports the relative water content (RWC) in the leaves of 13 woody species (between 77 and 91% of water).
But what about the rest of the tree. Of course I am not expecting an absolute answer as this would vary with species, age, season, health etc. But is there any study that has measured water content in whole trees? 
Relevant answer
Answer
Thanks Tim, 
This paper does look interesting, I will see if I can find more on their website. Thanks also for explaining the turgor loss point, makes more sense to me now.
Cheers,
Stephane.
  • asked a question related to Molecular Ecology
Question
6 answers
Neutral markers provide insight into levels of variation but do not take into account selective environmental pressures of a species and therefore  do not provide information in an evolutionary context as would the use of non-neutral markers. 
Is there any key significance or insight in using a neutral and non-neutral marker in conjunction for a population genetic study? 
Relevant answer
Answer
To begin with, it must be said that you may have some a priori idea about whether a given marker is neutral or has been under selection, however, you cannot know for sure. A coding locus with alleles responsible for different phenotypes could be neutral (phenotypes have or had the same fitness). A microsatellite or other "junk" DNA could be linked to a locus under selection and hitch-hike or have an unknown function and be responsible for changes in fitness. So beware of your assumptions about using a priori neutral and non-neutral markers.
Current widely accepted approach is to consider that the vast majority of the genome behaves neutral and only a handful of loci have been under recent selection. Therefore, we can perform demographic inference from lots of (randomly selected along the genome) markers. If some markers under selection are included, it is assumed that their signal is "crushed" compared to the main neutral signal carried by the overwhelming majority of neutral markers. See however Ewing & Jensen (2016, doi:10.1111/mec.13390).
Once the demographic history of the populations under study has been inferred one can try to detect loci that present diversity pattern incompatible to that history. These can be assumed to have been under selection.
  • asked a question related to Molecular Ecology
Question
4 answers
Can biochemical tests define microorganisms upto species level? Which manuals/ identification keys are used for identifying bacteria and fungus?
Relevant answer
Answer
Biochemical methods tests classify them to certain extent, which is just part of identification. You need molecular and advanced molecular methods to identify & classify them. Read below PDF....you would get some idea.
  • asked a question related to Molecular Ecology
Question
3 answers
Detecting different species of Oidium.
Relevant answer
Answer
Thanks your answer.
  • asked a question related to Molecular Ecology
Question
3 answers
Our lab uses ddRAD (a variant from the family of RAD-seq methods) for simultaneous SNP detection and genotyping. To find homologous loci across multiple individuals, we first cluster reads within an individual, based on some identity threshold (e.g. 90% similarity = same locus), and then do the same among all individuals in order to build a catalog of homologous sites across all individuals, again using some identity threshold. We do this using the program Vsearch (a centroid-clustering algorithm) as implemented by the pipeline pyRAD, but other popular RAD-seq assembling pipelines use a similar methodology (e.g. STACKS). 
My question is: How do we determine the "optimum" identity threshold for clustering? The issue is that if we have a threshold which is too strict, we erroneously split loci, but if our threshold is too relaxed we over-merge loci (for example clumping paralogs together). 
We have put some thought into this in our lab by looking at the relationship between various descriptors of our dataset and varying clustering thresholds (e.g. within-cluster genetic distance, number of parsimonious informative sites, number of loci failing paralog detection filtering, etc) but I am still having difficulty coming up with a logic for choosing a threshold. 
And of course this could also be done by considering the phylogenetic context- e.g. using timescale and mutation rate to predict average genetic distance within a locus, but this involves making a priori assumptions. 
Relevant answer
Answer
Hi Tyler, 
You might be interested in the papers by Mastretta-Yanes et al (2015) in Molecular Ecology (http://onlinelibrary.wiley.com/doi/10.1111/1755-0998.12291/abstract), and Viricel et al (2014) in Molecular Ecology Resources (http://onlinelibrary.wiley.com/doi/10.1111/1755-0998.12206/abstract)
Cheers, eric
  • asked a question related to Molecular Ecology
Question
5 answers
NMDS or PCA are usually employed for visualization of a (complex) beta diversity matrix. However, PCA try to maximize variation in several "ORTHOGONAL" axes (they should be independent between them).
When a factor has several groups, PCO (e.g. samples from 17 different environments) is not able to separate into the same number of clusters (even when statistics have a strong effect and high correlation).
Is there a way to visualize those complex data? Residuals? Discriminant Analyses?
Thanks
Relevant answer
HI,
My templates were bacterial DNA from bird eggshells.
I am working in your suggestions.
Thanks
  • asked a question related to Molecular Ecology
Question
16 answers
Hi everyone. I'm a sensory ecologist and I'm starting to get interested in ultraviolet light (also polarized light). I understand that birds and some insets (e.g. moths and butterflies) can sexually advertise in ultraviolet wavelengths. Still, there are so many diverse taxa of animals that can see UV that it seems like there have to be some more diverse uses for it other than prey identification and sexual advertising. I'm looking to create a bit of library of things that are strong reflectors of UV light and I'd really appreciate any you can contribute as mentions or more detailed references. Thanks so much.
Relevant answer
Answer
Urine can be "seen" in the UV helping birds of prey find rodents. Berries differ in their UV reflectance. Some example images of flowers and berries are available at http://www.uv4plants.org/gallery/plants-and-flowers-in-vis-and-uv-images/ as pairs of visible and UV images. Images are from Lasse Ylianttila.
Below are links to some publications I am aware of, but my expertise is more on plants than on animal vision.
  • asked a question related to Molecular Ecology
Question
1 answer
When it's running it looks all good, like I'm going to get good sequence. Then it just puts out this small file that only contains the RAW data and I'm not able to perform base calling on it. I think it is related to the sample name because all the ones that fail have the same beginning in their sample name; "BPI". That said, these sample have worked before no problem, so I am unsure what has changed. I've had trouble with the sample sheets before, they weren't loading properly because of something my computer does to them. Now I just copy paste the sample names into a sheet on another computer. This happens on both our 3130 and 3730. Has anyone else seen this before? Thanks so much for your time!
Relevant answer
Answer
SOLVED. There was a non-breaking space in my sample names. Simply removed that and all is well again.
  • asked a question related to Molecular Ecology
Question
8 answers
 In the research paper I am reading, they used ND3 gene and cyt-b gene to be markers for studying bird populations but used COI gene for parasitic insects, so I was wondering what the criteria for choosing the markers are. 
Relevant answer
Answer
Dear Mr. Phuangphong
In my experience, choosing markers depends on the questions you are planning to answer, and also on the sequence resources already available for comparison. In the case of using different mtDNA regions, as you pointed, usually the choice is based on te available sequences in the databases. COI has been extensively used for insects, and therefore was selected as the region for barcoding especially these organisms. On the other hand, cyt-b was historically used for mammals, and therefore, the database for cyt-B has more data than the COI database for mammals. I am not a specialist in birds, but I would guess that cyt-B and ND3 might have a good database for these organisms.
Hope this helps,
Best
  • asked a question related to Molecular Ecology
Question
5 answers
I work on an ancient protein family that exists in both eukaryotes and prokaryotes and my interest is in elucidating its deeper branches. When I pull down the data from NCBI I get about 6000 sequences. If I use an algorithm to help me cluster similar proteins (e.g. 80% similar) it gets reduced, and if I do my clustering with 50% similarity I get about 550.
The question is, in your experience, which one would you choose:
- use the smaller dataset (or even smaller) and do the most sophisticated, yet computationally intensive, analyses you can hoping that if there is a signal, these analyses would pick it up and thus the deeper branches get better support.
- use a lot of data and hope that then the intermediate evolutionary steps could be inferred more easily and the deeper branches get more support?
Relevant answer
Answer
The 2nd option is probably more likely to give a quality answer. If you have the time/computing power I think you should look at the larger dataset, too. If your clustering combines particularly disparate sequences you may be thinning out your taxa distribution and inviting more opportunity for long branch attraction. I would consider comparing the topologies and look for places where lots of relative diversity was collapsed in one area but not in other areas of the tree. If it is even throughout, the smaller dataset is probably much more likely to give a fair answer.
  • asked a question related to Molecular Ecology
Question
6 answers
Hi all,
I have no experience with CARD-FISH. But right now I suffered a lot from the autofluorescence of the sponges symbionts, which I tried to target. The autofluorescence was strong and universal. What's worse, the de-fluorescence methods didn't work well. So I am thinking about CARD-FISH.
My samples were collected in June 2013. Some of them were immediately fixed by 4% paraformaldehyde while some of them were fixed by RNA later. All samples were restored under -20/80 centigrade. From my own opinion, none of these samples should be used for CRAD-FISH now. 
Any input is welcome.
Cheers! Fang
Relevant answer
Answer
if it's just stored in PBS without fixation, probably not suitable for FISH
  • asked a question related to Molecular Ecology
Question
4 answers
Can anyone share how to study or anlyze the crossability barriers in wide hybridization crosses please?
Any vegetable crop...thank you
  • asked a question related to Molecular Ecology
Question
10 answers
The genus Cebuella in Colombia has been replaced to Callitrix and i have not found the right explanation. 
I cannot understand why the taxonomist consider the genus Cebuella in Colombia as Callitrix nowdays... I have been told there is a mollecular evidence somewhere.. i am not sure whether it is Tagliaro et al. 2000.
Many thanks in advance.
Relevant answer
Answer
Dear Deborah,
Whether you call these animals Cebuella pygmaeus or Callitrix pygmaea it still remains the same species! Morphology AND molecules seem to indicate that this species is a separated entity within the clade with species of Callitrix. If this is enough to consider it a member of a separate genus is completely dependent of the opinion of the taxonomist: there are no "rules" for this. So you are perfectly entitled to call this species Cebuella pygmaeus (and this might even be the most pragmatic solution). One remark however: do NOT speak about Cebuella when you mean the species; in other words: do not use the name of the genus when you want to indicate a species.
So Johana, you have the choice between the names Cebuella pygmaeus or Callitrix pygmaea for the species that was described for the first time by Spix in 1823 as Iacchys pygmaeus. Because the genus name Iacchus has not been used since, it can be considered a "nomen oblitum" (but the species is valid).
  • asked a question related to Molecular Ecology
Question
6 answers
The tissue of the non-fleshy corals such as Acroporidae, and Pocilloporidae family contains much skeletons than the tissues. I had been saw my colleagues working on coral histology and they use 7-10% formic acid to decalcify the skeletons to get the tissue to make histological slide, but is it also safe for preserving genetic materials on corals? If it's not safe, how could I harvest large amount of coral tissue for genetic analyses from non-fleshy corals?
Relevant answer
Answer
Hi Stevanus,
I found the best method for myself (currently working on extracting DNA from a number of scleractinian families) is to crush the required amount of coral with a mortar and pestle. The skeleton does a good job in lysing the tissue. I usually then store the crushed tissue in a guanidium thiocynate  based "CHAOS" buffer for a couple days, followed by a ProK digestion then a standard kit or PCI cleanup... I get very high quality and purity DNA using this method.
If you have coral samples stored in ETOH, you will have to wash the sample thoroughly in distilled water. If you vortex your sample in distilled water, it also helps remove most of the mucus that the coral produces. Mucus can be a problem, especially as it can block silica based flow through tubes and can inhibit PCR.
Good luck!
  • asked a question related to Molecular Ecology
Question
4 answers
When dealing with molecular dating, ones referring to a reference time based on fossil records. How feasible/comparable can these data be applied to study the evolution of a particular genus when we can assessed to some of the widely used genetic markers, e.g., ribosomal DNA (28S or LSU, 18S or SSU), nuclear encoded gene (ITS1-5.8S-ITS2), mitochondrial DNA (cox1) and plastid gene (RUBISCO or rbcL)?
Since each of the gene markers evolve independently with different evolutionary rate, would the calibration be optimal based on only one gene tree? Which gene marker is more preferable and what kind of criteria should we follow? Should we combine the gene trees follow by the calibration? Or, concatenate the sequence data of all the available gene markers, generate a tree, only then calibration?
Thanks,
HongChang
Relevant answer
Answer
BEAST software should be useful for you. It considers each gene information (seperate clock models, substituon model etc.) at the same time and it also handle with incongruence among gene trees. 
  • asked a question related to Molecular Ecology
Question
9 answers
After some difficulties, working around security features, I have managed to download and install the source code. Now I am having difficulty getting it to read my data. I get the error "Cannot open data file "<filename>"
If it is being used by another application, close it first, then press RETURN
Otherwise enter a new name for the data file
Press ctrl+c if you wish to stop the program now:
Has anyone else had this problem? Did you manage to solve it?
Relevant answer
Answer
Please be aware that "sudo" means that the software has root = complete access to your computer and private data. It is a command for administrators and should be used only in such situations. 
  • asked a question related to Molecular Ecology
Question
6 answers
Hi all. I'm not an evolutionary expert. I want to test if there is any evolutionary relationship between three enzyme families (two of them are known and the other is the one that I'm interesting in). When I make a quick phylogenetic analysis, the two known families group separately, and main group with one of them. The point is that I think that maybe there's a relation between all of them. How can I test that? I think an outer group should be the best way; but I don't know how to choose the correct one. Any suggestions?
Lucio
Relevant answer
Answer
I agree heavily with Jose Tomas Matus that using CDD to find families with matching domains is a very good idea. I would just caution that you may need to be careful about what you get from this search. If you pull a divergent protein where this domain is only a small part of the overall sequence you will wind up aligning only that domain and your tree will be a history of said region. This may not be a problem depending upon your circumstances, but it might be a major issue that gives you a false result.
One way of augmenting a CDD search is through a set of BLAST searches. I assume you already have accepted sequences for each of your families, plus your own sequences. Use a number of each from all three groups as seeds for BLAST searches and take a large number of hits (preferably, mostly those you know are not part of your groups). Ideally, these will coincide with your CDD searches and you can include a significant number of them to provide shape to your tree. A large and varied outgroup set can help ameliorate some of the problems of using a single too-distant or too-close taxon. If your new group falls in or sister to the two know groups with no outgroups among or between, then you have some good evidence that your 3rd group is closely related to the knowns.
  • asked a question related to Molecular Ecology
Question
6 answers
I am working with RAD-sequences and trying to build phylogeny of the genus, but I realized that usual tree-building methods are not made for complex reticulated history. In contrary, such methods like Bayesian clustering in Structure, DAPC in adegenet or Patterson D-statistics do not reconstruct phylogeny and/or can estimate introgression in the the known hybrid zones/hybridizing pairs of taxa.
Relevant answer
Answer
I think that the method that you use here really depends upon the question that you want to answer. A bifurcating phylogenetic tree applied to snp data can still be a very useful thing if one wants to establish relationships between a closely related group of individuals. But I think its important to acknowledge the limitations of the resulting toplogy. In reality, every gene will have a different history, but in most cases there will probably be an 'average' signal when one considers many loci. For example, this average might reflect the history of geographic divergence between a group of closely related taxa. But say that following divergence, introgression lead to the sharing of some genomic regions that were favored in some or all of the taxa. Loci like these will have very different geneaologies which will be swamped by the average signal in a standard phylogenetic analysis. 
If you do want to make an attempt at inferring introgression events in a tree-based analysis, you could check out pyrad (http://dereneaton.com/software/pyrad/), or Treemix, from the Pritchard lab. 
  • asked a question related to Molecular Ecology
Question
6 answers
Unlike Trachelomonas superba, which is furnished with spines all throughout this one have  spines on two sides only.
Thanks in advance.!
Relevant answer
Answer
T. hispida var. duplex Deflandre 1926? There are experimental work done on the degree of mineralization in Trachelomonas, may be 30+ years ago. This can depend upon environmental factors..... like chelation and availability of iron and manganese (redox)
  • asked a question related to Molecular Ecology
Question
5 answers
Hey everyone.
Has anyone read the Mol Ecol paper by Ferretti et al. 2013: Population genomics from pool sequencing? Specifically, has anyone tried to use their calculation of Fst from estimated nucleotide diversity?
For some reason, what ever I do I keep getting an Fst of zero -- even for very simple data with only two populations and very different nucleotide diversities. I have attached the equations, a description of the parameters, and my calculations to this question.
It could be that I have misread something important, or my order of operations in the summations is wrong.
For pooled data, being able to use nucleotide diversity to calculate Fst would be such a huge advantage to any study. Can anyone see what I have done wrong?
Relevant answer
Answer
Sebastian and Xavier are right. Example: Sequences from two (haploid) populations
pop1:
ACAAAT
ACAAGT
pop2:
CCAAGT
CCAAGT
In this case, the internal variability Hs is 1/6=0.167 for pop1 and 0 for pop2. However, the average pairwise nucleotide diversity between the populations is 0.25 because of the fixed difference A/C between populations in the first base.
  • asked a question related to Molecular Ecology
Question
5 answers
Hi everyone. I've been looking for a software in which I can simulate hybrid crosses between potential parental genotypes. I've found that HybridLab software (Molecular Ecology Notes (2006) 6, 971–973 doi: 10.1111/j.1471-8286.2006.01433.x) could be useful. However the link provided by the authors seems to be broken. Does anyone know where should i go to download this software? Is there any other software I could use?
Any help will be appreciated :)
Relevant answer
Answer
I have it if you still need it.
  • asked a question related to Molecular Ecology
Question
8 answers
The internal transcribed spacer regions (ITS1-5.8S-ITS2) is amplifying and sequencing beautifully on the stock DNA extraction from my target plant species (member of the Fabaceae).
But none of the chloroplast regions I use amplify. I have used stock, -1 (i.e. 10^-1), -2 and -3 dilutions. Positives from other species amplify - so not a PCR protocol or primer problem. 
The only thing I can think of is that there is a large disparity between the chloroplast and nuclear DNA amounts. My feeling is that there should be more chloroplast copies than nuclear, but surely the -3 dilution should be sufficient?
My concern is that there may be too few cpDNA copies relative to the nDNA and that I should be using more stock (I am using a new extraction kit on a new target species).
(Just a note - I have no access to a Nanodrop, and the Qubit is out of operation at present).
Any thoughts or suggestions would be greatly appreciated.
Relevant answer
Answer
Check 1 ul of your DNA in an agarose gel (0.8%) made with TAE.
If you can not see  a high molecular weight band, change your DNA extraction kit. (or check with the company providing the kit).
DNA from members of the Fabaceae should extract nicely....If you perform your ITS PCRs with DNA from the stock, then something is going wrong with the extraction
  • asked a question related to Molecular Ecology
Question
13 answers
Hello every body
I am working in a project involving screening of spiny lobster using 100 microsatellite groped into multiplexes.. During my research, I faced some markers that succeed in some samples but not amplified in others. I am sure it is not due to experimental mistakes since I manage my protocol to be uniform with all samples.. I repeat it several time with same observations. I sequenced the amplified PCR products and Blast it. I found some similarity with part of the mRNA in other species.. I found some literature giving reasons of this phenomena in cross species amplification. But my observation is in the same species from the same geographical area.
Relevant answer
Answer
Null alleles is a likely answer, and some samples will never amplify because of that (but as Roberta suggests, this might not be a major problem with analysis). If you decide to try one more amplification, you might try spiking the PCR of the reluctant samples with magnesium chloride. We discovered this helped some samples significantly - I would try increasing by just 1 mM and see if you get results... just be sure you are not getting non-specific priming... microsatellites are nice because you know where the alleles should be - if you get peaks/bands outside of that region with extra MgCl, back down on how much you are putting in...
Good luck!
  • asked a question related to Molecular Ecology
Question
3 answers
I am working on re-designing 96 barcode adapters for RAD-seq (or GBS or genotyping-by-sequencing or ddRAD or whatever you prefer to call it) for our lab.  I would think that adapters that would work for a PstI overhang would also work for an NsiI overhang, since the overhang is the same.  However, I have noticed that the conventional wisdom is not to recreate the restriction cut site, which is why none of the standard barcode sets for PstI end in a C.  By that logic, if I was going to use the barcodes for NsiI I would not want any barcodes to end in an A either.  But if I heat-inactivate the NsiI before the ligation step, why does it matter if the ligation re-creates the cut site?  Is there a bioinformatics problem, or is it just the wet lab problem of residual restriction enzyme un-doing the ligation?
Relevant answer
Answer
Some people also like to digest again at a later stage to remove chimeric fragments that ligated together at the cut sites. 
  • asked a question related to Molecular Ecology
Question
3 answers
I am looking for fresh samples of serpulid genus Hydroides fixed in ethanol. I have this unique opportunity to sequence all Hydroides we can get to compile a barcoding database for this important fouling and invasive marine taxon. As an incentive, I can offer a co-authorship (depending on the amount of the material) or  can send thee very early version of the "Hydroides of the World" book fully illustrated by original photos.
Relevant answer
Answer
Yes, it is H. norvegicus. In the case of potential neotypification these specimens should be of more interest than the previous as they are somewhat closer to the (potential) type locality. Although this is not too precise in Gunnerus' original description I would believe so. 
Cheers, Torkild
  • asked a question related to Molecular Ecology
Question
6 answers
Does anyone know of any papers that discuss problems associated with a lack of source populations when attempting to correctly quantify introgression with programs such as STRUCTURE, BAPS or Flock? I'm working on a simulation study that suggests that without both source populations admixture levels are falsely low (if run only with one parental) or falsely high (if run with one parental and a different source pop as a "surrogate"). Trying to wrap my head around this, and wondering if anyone else has experienced this problem. Also wondering if anyone has shown STRUCTURE's inability to correctly quantify ancestry in this scenario in the past. Thanks!
Relevant answer
Answer
Dear Spencer, we have been doing some structure and admixture analysis on wild species populations, using also STRUCTURE and NEWHYBRIDS (among other approaches). At least in two cases (different species and scenarios), STRUCTURE identified, as different genetic clusters, two populations we suspect that correspond to a parental and an admixed population (in the absence of the second parental population). I say that we suspect that because we run the same pairs of populations in NEWHYBRIDS and we got  very good agreement with the two clusters identified in STRUCTURE and the two identified in NEWHYBRIDS. The interesting thing is that NEWHYBRIDS identified individuals from one of the populations as parental, and the individuals from the second as second generation hybrids. In both cases, we did not included (and were not identified) individuals from the second parental population. These results seem to make sense on the biogeographic context. In one case, we suppose that the second parental population is actually a source population from a founder event and that the admixed population is actually resulting from the admixture between propagule population and the native (pre-existing) parental population in the arrival place. On the other case, we have one population from one side of a mountain range and other in the mountain range (but not on the other side). We suspected that mountain population should be receiving genes from both sides and STRUCTURE identified it as an independent genetic cluster and NEWHYBRIDS identified it as an hybrid cluster.
So, trying to answer your question, I think that (in the absence of one of the parental populations), STRUCTURE is able to identify admixed population as a different cluster, but might provide no idea about the nature (admixed/no admixed) of that population. However, if you test the pair (putative parental + putative admixed) in NEWHYBRIDS, you might be able to detect if individuals are actually of admixed origin or not. I think this happens because NEWHYBRIDS algorithm explicitly searches for hybrid and parental classes and allways assume two parental classes (that it will try to reconstruct based on a bayesian algorithm). Even if you have no individuals from a second parental population, it might be able to reconstruct the hypothetic non-sampled parental population. At least with us, it seems to be doing that, at least in two different examples, and on a consistent way. Despite we only provide the genotypes to both programs, without any a priori information, it ends up consistently with the same outcomes.
From the scenarios I've already analysed with STRUCTURE, I found that when you have both parental and an admixed population, STRUCTURE detects the admixed population as a "gradient" of individuals with different proportions of genome assigned to each one of the two parental populations.
Well, I am not sure if this helps! If not, please apologise me for the long argument...
  • asked a question related to Molecular Ecology
Question
1 answer
What is the interpretation of Mendelbrot model in vegetation data, when I use the function "radfit" in Vegan, part of my data has Mendelbrot model? And I can't find a good article that describes the interpretation of this model.
Relevant answer
Answer
See the attached files.  It's one of several models  for rank abundance data against which data are often fit. Interpretation is complex, with arguments that provides evidence for neutral & niche based community structure. 
  • asked a question related to Molecular Ecology
Question
4 answers
It can be a naive question, but... If a historical expansion event happens, which is evident in mtDNA sequences, is it possible that expansion might have obscured  signals from a past divergence event? If so, should we see this obscuring effect in both nuclear and mtDNA markers to the same extent? Or do demographic changes affect mtDNA more dramatically compared to nuclear DNA because mtDNA has a smaller effective population size? More specifically, is it possible that nuDNA maintains the signal from past divergence event whereas mtDNA loses it after a population expansion event?
Thank you.
Relevant answer
Answer
Hi Semir,
Thank you for your answer.
  • asked a question related to Molecular Ecology
Question
3 answers
I would like to select a molecular marker to compare about 100 strains of Trichoderma harzianum. I think these isolates be too similar, so an appropriate  maker need to differentiate them.  
Relevant answer
Answer
Since you have one species of Trichoderma only. The best marker will be SSRs. Don't do simple diversity but see the population structure which will help you in indentifying the various evolutionary forces shaping the fungus tragteroies
  • asked a question related to Molecular Ecology
Question
5 answers
I'm working with mtDNA control region in parrots and have been trying to understand exactly how to identify or check sequence data for numts?  So far I've found suggestions to check sequence data against others possibly published/genbank, and to use diluted DNA, but I wonder if there are anymore suggestions out there? Many thanks everyone.
Relevant answer
Answer
If you are having difficulty distinguishing them from the outset and have access to some soft tissue (liver is ideal), you can isolate mtDNA by differential centrifugation, then redesign primers to be specific to that sequence. As numts have a much lower mutation rate, and there is usually only one amplifying, they are usually easy to identify as a contaminating sequence. They are often useful in themselves as an outgroup for the real mtDNA- the low substitution rate means that you almost have the equivalent of a fossil ancestral genome to work with. So you might want to design primers for the numt itself too.
  • asked a question related to Molecular Ecology
Question
2 answers
I am conducting a microbiome analysis of Anolis lizards using 16s microbial primers. I have dissected the fecal chambers of the GI tracts to remove fecal material (without host tissue) but some samples have large pieces of exoskeleton. I am concerned this may bias the microbial sequencing results as I am conducting both chemical and mechanical lysing in my extraction protocol that will most likely incorporate prey microbial DNA from these exoskeletons into my extractions. A colleague of mine has done a similar study of passerines, and noticed the fecal samples significantly differed in microbiome from oral and cloacal swabs of the same individuals collected at the same time. It seems the prey tissue may affect the microbial assessment.
Has anyone tried to eliminate this potential bias? This caveat has made me think more about "host microbiome" in general. While prey preference has been shown to influence microbial assemblages in hosts, how do we know if this is because the prey microbes have been incorporated into the host holobiome or simply because it is temporarily present during the analyses?
Relevant answer
Answer
This is a great question and I'm not sure that there's a perfect answer.  I have not worked with samples from poikilotherms but we have a similar issue in mammalian samples, i.e., determining which sequence data represent taxa patently colonizing the GI tract of the host in question and which represent DNA present in the food.  With untreated rodent chow, much of this residual DNA may represent spores or viable organisms but, even in our analyses of autoclaved, irradiated food (fed in strict barrier and germ-free facilities), we detect dozens of unexpected taxa.  The DNA is presumably from dead bacteria, killed during autoclaving and irradiation, but it does introduce a confounding factor in some situations.  I'm not aware of a good method of identifying and "subtracting" those food-associated sequences from the data matrix and, in general, I leave them in the data and interpret everything based on the relative abundance of suspect hits and what is known about those microbes in question.  As an example of the former, there are microbes such as segmented filamentous bacteria (SFB) which have significant effects on host physiology and are of interest in many mouse disease models.  However, it's also a spore-former and common contaminant of rodent chow.  If I detect SFB in a mouse fecal sample, I check to see whether I have 1 or 2 sequences, or 1 or 2 thousand sequences.  As a second example, we frequently detect sequences that annotate to "order Streptophyta" - these are plant-associated microbes so I assume (right or wrong) that this is not patent colonization.  I was heartened to see sequence data from germ-free mouse feces was largely Order Streptophyta.  As I said before, I'm not sure if there's a perfect answer although there may be some advance informatics approaches to filtering out those OTUs of which I'm just not aware.
Hope this helps and good luck.  Aaron
  • asked a question related to Molecular Ecology
Question
5 answers
I have a dataset of COI sequences and I'd like to obtain Bayesian Skyline Plots (BSPs) with BEAST for my populations. I made 5 replicates runs obtaining 5 .log and 5 .trees files. I used LogCombiner 2.2.0 to obtain single .log and .trees files from the 5 replicates, in order to construct the BSPs with Tracer. LogCombiner was able to construct a combo file for trees, but it did not for .log files. The program stops without producing any file and without giving any error message. Actually it made nothing....! Any suggestion or hint?
Relevant answer
Answer
Thanks everybody! LogCombiner seems to work when I use an old version (1.7).
  • asked a question related to Molecular Ecology
Question
5 answers
Using admixture model as is, we have enough strong signal for some individuals from each locality and weak signal for others. And we want to use admixture-locprior algorithm to test may these individuals belong to known populations, too, or they may originate from additional populations located outside of the all locations in studied area.
Thank you.
Relevant answer