Science topic

Molecular Evolution - Science topic

The process of cumulative change at the level of DNA, RNA, and PROTEINS, over successive generations.
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I am try to find out the main differences between haplotype and DTE analyses. To my knowledge, the outer Haplotypes are newly created, while the inner ones are evolutionary older than the outers. What happen if we find out different results in DTE?
I mean I found that some taxa are evolutionary young according to DTE, while haplotype analysis showed me the opposite results.
Would you please let me know, how I must interpret this difference?
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Dear Atena,
You should bare in mind that relationships among taxa in haplotype networks are not necessarily equivalent to their relationships in phylogenetic trees (such as Bayesian, maximum likelihood or parsimony). This is because a haplotype network is usually calculated based on a distance matrix, thus it clusters individuals according to their genetic distances. This may or may not reflect the 'true' phylogeny.
So the short answer is that the two are not easily comparable.
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The origin of life on Earth is still not known. Some maintain that rather than it having originated here as a result of complex organic chemical reactions that it arrived fully formed from space on comets.
What evidence is there that would support this claim?
Even if true it would still not fully explain how life came about in the universe. Does anyone have any suggestions as to that conundrum?
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The origin of life is not a singularity, so, it is also possible. Organic molecules might be developed in cometary tails, and in fullerene cages in interstellar dust, nebulae, comets, etc. Fullerenes are likely UV resistant and can wrap and protect organic molecules. They can also keep water on the cage surface. This is mostly related to RNA world and PAH world hypothesises.
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I am looking for information about amino acid substitution models. I read this paper (Trends in substitution models of molecular evolution, doi: 10.3389/fgene.2015.00319) but I was wondering if there is any other review or book/book chapter recommended.
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Thank you!
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I want to use BEAST to do EBSP analyses with two loci. I open two "input.nex" files in BEAUti to generate a "output.xml" file (In the Trees panel select Extended Bayesian skyline plot for the tree prior), and then run BEAST. I do not know if this is right and I do not know what to do next. I can not construct the trend of demographic history in Tracer just like BSP. I got one log file but two trees files (for each locus), and I do not know how to import both tree files into Tracer.
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One of the best websites to find the answers to these questions is the following link:
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I am trying to find an appropriate selection test to evaluate whether selection in a gene was intensified after a specific time (i.e. I am using phylogenetic dating to estimate an evolutionary event). My hypothesis is that after the event the selection strength in the gene increased.
I have been searching for different tests to test this but I haven't found anything 100% convincing.
For example, branch tests implemented in PAML (codeML) can evaluate differences between specified foreground branches and background branches. Although, I am not sure if I should mark all the branches of the tree after the event.
Another alternative is using HyPhy, with a test like RELAX for detecting shifts along branches. I am also unsure if this is appropriate. BUSTED looks like a gene-wide average, similar to the codeML models. And aBSREL only takes into account positive selection.
As you can see I am very lost here and any help is very much appreciated.
Please, kindly advise.
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A molecular clock test will tell you if there is a change in the rate of evolution somewhere and will in some programs idicate which branches depart from equal rates.
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Hello all,
I have population genetics data (mtDNA) for 5 populations (n=20) of the same species. All neutrality tests (Fu's Fs, Tajima's D, Fu & Li's F, Fu & Li's D, Roza's R2, Strobeck's S) are non-significant so no signal of deviation from neutrality is there. However mismatch distribution analysis (with demographic expansion model) for 3 of the 2 populations are unimodal, the other two are multimodal. SSD for multimodal distributions are significant and the rest are not, suggesting three of the populations are not significantly different than population expansion model. Raggedness index for all of the populations are non-significant suggesting all of the populations are not significantly different than population expansion model.
1) What could cause the contrasting results from neutrality tests and mismatch analysis?
2) Why is raggedness index failing to capture the multi-modal distribution of the two populations while SSD can?
Another contradiction I face is between the actual mismatch distribution and the mismatch statistics. I attached a mismatch distribution for a different population. It is unimodal (albeti skewed) and observed values seem like a real good match to the simulated values. However SSD (0.00236) and raggedness index (0.07244) for this plot are both significant.
I am a bit lost in what to deduce from these results. Can you help me please?
Thanks!
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Can anybody help me how can I interpret my opposite results between neutrality test-mismatch curve and SSD-raggedness index (Hri) tests?
In my results, insignificant Tajima’s D= -0.76 (P= 0.24); Fu’s Fs= -1.59 (P= 0.27) and multi modal observed MMD curve denotes populations are at equilibrium. However, SSD = 0.016 (P = 0.48) and Hri = 0.009 (P = 0.15) are insignificant which denotes population expansion.
Please see the attached picture. I used Arlequin program for analysis. In my d-loop DNA sequence, I have long indel zones in the sequences.
Can anybody help me, please?
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Hi everyone,
I am having some ideas to conduct a set of analyses concerning insect speciation and phenotypic diversification. For this, I will need a species-level phylogeny with complete taxon sampling (or as close as possible to this) - that is, few known species will be lacking in the tree. ANY group of insects is fine. I also need the phylogeny to be as big as possible, say at least 80-90 terminal taxa. Any thoughts?
ps. I also welcome collaborators, please contact if you like these topics :)
bruno
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Hi Bruno. This is really not the case. I'll check if there is something for Cicadidae but I don't think so. Cheers!
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Hi All,
Are you an expertise in getting ancient DNA? Currently I am looking researchers who have expertise in getting DNA from old herbaria collections. Even better if you have it with Ascomycota (Fungi). If so and you like to collaborate with taxonomist to work in projects about systematics, evolution and biogeography, please contact me at lquijull@gmail.com or luis_quijada@fas.harvard.edu and write in the subject of the email "ancient DNA". I am looking for collaboration to learn these technics but also you will be coauthor of the results of this project that start in July.
Best wishes
Luis
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we have extracted DNA from ca. 30-50 years old herbarium basidio samples including some types, though this age is not that “ancient”…
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Who gives the last word about the evolutionary process, genetics or ecology?
In other words, are ecological interactions driven by any genetic phenomenon? Or is it genetics that has been molded by ecology?
[I’m a Brazilian biologist and writer. I write about science – I have just released a new book, O que é darwinismo (What is Darwinism, in Portuguese) – and would like to know the opinion of colleagues from other countries (from any field of scientific knowledge).]
See also What do you think about fitness, adaptation and natural selection? (https://www.researchgate.net/post/What_do_you_think_about_fitness_adaptation_and_natural_selection)
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Nobel Prize winner Niko Tinbergen observed in his now classic 1963 paper, "On the aims and methods of ethology" (Zeitschrift fur Tierpsychologie 20: 410-433) that in order to adequately analyze observed patterns of behavioural ecology in a species, it is necessary to distinguish between *ultimate* and *proximate* causative factors. Ultimate factors include: i) the *function* (or "adaptive value") of a behaviour, and ii) the "phylogeny" (or evolutionary history) of a behaviour; proximate factors include: i) *ontogeny* (or behavioural changes related tp growth and development), and ii)*proximate conditions* (i.e., that which has happened in the recent past and that which is going on under current ecological conditions). So, what is needed in trying to gain insight on biological evolution is an holistic perspective that incorporates *both* genetics and ecology. A perfect example of the value of this integrated approach is the need for up-to-date data in both the genetic and ecological realms in order to deal with conservation biology issues.
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Dear professors
Respectfully, i know that sequences can help us to reveal the most common ancestor in phylogenetic studies. However, in genetic diversity studies using IRAP, SSR, markers etc, i want to know if it is possible to estimate Tmarca? If the answer is yes, How? which software can help us?
I will appreciate if anyone can help me.
Regards
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Dear @Atena Eslami
Quite a while ago I did my PhD on evolutionary taxonomy of a family of fish (Platycephalidae) using allozyme markers. I tested various methods of coding these markers and found that a binary coding system proved to be the best. Details can be found in:
Phylogeny of Australian species of flatheads (Teleostei, Platycephalidae) as determined by allozyme electrophoresis
CP Keenan - Journal of Fish Biology, 1991, Vol39: 237-249
I hope this helps.
Cheers, Clive Keenan
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Can someone provide suggestions on which methods / softwares can perform fast and reliable estimates of diversification rates (speciation - extinction) using very large phylogenetic trees (> 10 000 tips)? 
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BAMM, LASER, and APE
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I am working with some strains of cyanobacteria isolated from Zimbabwe's Manyame catchment - I want to extract their genomic material so that I may use it to confirm their identity. In my lab I do not have the capacity to buy DNA extraction kits so I have to use other means. I tried to use the CTAB method but failing to get good results. May I have some options which I can try with my samples.
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I know that this topic is kinda controversial to some, but, thru discussion, i suppose we can achieve greater understanding of the topic.
So, I wanna know what are the aspects, process, evidence, etc about evolution that you think is problematic?
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Hi everyone,
There seems to be a fundamental misunderstanding here. Evolution is by no means problematic, unscientific or lacks sufficient evidence. Speaking of the fossil record and that organisms don’t change – they do indeed and they do so quite dramatically and over short and long time periods. As a paleontologist, I study exactly those changes over millions of years. I do not want to criticize your belief, for I do not see any problem whatsoever to combine religion and science, they are just very different views on the subject. But please do not mix up a scientific approach, which tries to objectively explain how life has evolved based on facts, and a religious interpretation. Of course many findings of science are not 100% certain and it also involves some interpretation, but the same is true for every field of science.
Best regards, Thomas
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I would like to analyze adaptive molecular evolution of a insect mitochondrial gene. Could anyone please suggest me the procedure or tools to perform this.
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Madhav, That's just fine. But there is so much more to testing for selection than simple dn/ds. Of course that is exactly where to start and I usually do a dn/ds with Seaview where I also do the alignment and maximum likelihood.
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I want to analyze the association/correlation between the rate of molecular evolution (dS, dN, omega) and different quantitative traits (eg. temperature).
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I second what was said above. A mantel test should work. Because temperature and latitude are correlated, you can also use a partial mantel test.
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Citations that appear in the BODY of published papers contribute to metrics (e.g. h-index) that account for the impact of the work of a particular researcher. I acknowledge that the suitability of these indexes is debatable. In the meantime, I wonder why the citations that appear in Supporting Information (particularly those that refers to relevant methods and databases that were used in the papers) are not included in the calculation of such indexes.
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Yes! Hope journals will/are considering this issue.
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Good day house. I am working on Molecular Evolution and Phylogenetics of some organisms. Before conducting my Synonymous to Non Synonymous substitutions after Sanger Sequencing, I would need to get various statistics from my DNA sequence. Asides these:
1. Base Composition
2. GC content
3. Word pattern
4. Over represented and under represented words
What are the other DNA Sequence statistics that could/might bring out something from the sequence with respect to analysis?
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It depends a bit on what type of organisms you are studying: bacteria, virus, plant, animal, etc. For some organisms it is interesting to look at CpG dinucleotide because this gets methylated and then mutates to ApG or CpT so genomes with meythylation are very much depleted in CpG dinucleotide content. For bacteria it can be interesting to look for transposons and other mobile elements. For bacterial genomes the "GC skew" can be useful in identifying genes and direction of replication.
It can be very informative to do most of these analyses in a sliding window, rather than over the entire sequence. I am attaching a figure made using the gwBrowser genome browser which plots many DNA attributes or statistics as rings. In the pink vs cyan inner ring you see the GCskew and how it correlates with the gene orientation on the plasmid (Blue vs Red on the outermost ring). The dogma is that GC skew is due to direction of replication and not direction of transcription. But my studies of Clostridia make me question the dogma.
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Oldest Species in the World and Cancer?
Cancer being a vulnerability to mutation deeply rooted in our DNA.
Sharks, crocodiles, horseshoe crabs.. seems like if they stood up to the test of time very well.
I tend to think that they could have evolved mechanisms to have reduced burdens of cancer having been around so long.
These species have been around for a while. Looking at some of the fossil records they have changed little in terms of structure.
So they have been very successful for a long time.
Could evolution of their genetic stability give them endurance to stand up to so much time and still be successful with such little change?
"Together we will"
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The Phylum Onychophora are a very small group of terrestrial animals, commonly known as peripatus or velvet worms. Onychophorans are considered “living fossils” because they have retained an evolutionary stage intermediate between a polychaete (legged marine worm) and a myriapod (group of terrestrial arthropods that includes centipedes) for 500 million years.
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Human self-medication behavior is normally considered as a learned behavior, even into a cultural context.
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Dear María,
¡Muchas gracias! This paper looks very useful to draft an answer to this complex question of the origins of self-medication behavior. I believe that animal self-medication is key to understand its evolution.
Recibe un saludo muy especial.
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Is there anything similar to a simulation that starts with a random genetic code and some genes that are translated into aaRS proteins which change the genetic code because the new aaRS aren't the same as the ones generated by the previous translation?
Is there literature about how the genetic code got stable? (stable means each codon has just one translation)
Thanks in advance.
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Actually, in my opinion, there is a substantial record on just the opposite - that the genetic code has still been evolving and is not stable. However, if analyzing these phenomena, one can conclude about how the genetic code has been conserved (at least to the degree it has been, at all)). It may concern in particular the phenomenon of codon bias; see: Sharp & Lee, Nucl. Acid Res. 1987; 15(3) 1281 (this one you must know well); Kudla et al., Science 324, 255 (2009); DOI: 10.1126/science.1170160; Buhr et al., Mol Cell. 2016 February 4; 61(3): 341–351. doi:10.1016/j.molcel.2016.01.00;
As to POLYSEMOUS (SIC!) codons, there is an interesting paper (quite an old one) showing that the genetic code is not set absolutely, and it keeps evolving: Suzuki et al., The EMBO Journal Vol.16 No.5 pp.1122–1134, 1997
I hope tnis is at least a little of what you need. I thing the codon bias (preference) is the main evolutionary force, in particular if you join it with the phenomenon of polysemous codons. A lot to think on, isn't it?
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Are there simulations that...
- simulate changes in the tRNA pool?
- simulate flexizymes?
- simulate aaRS?
I did a lot of research but didn't find practical approachs on the role of these three components in genetic code evolution (either mathematical or computer simulations or laboratory experiments).
I'd appreciate any information in this direction.
Thanks in advance.
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Hi Hanna!
If you are interested about more theoretical aspects of genetic code emergence, especially about a coevolving nucleic acid/protein world and gene-replicase-translatase systems, I suggest to take at look at the work of Peter R Wills. Good papers to start with are:
(about a theoretical model of self-organization of the coding system)
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When tRNAs can have three, four and five base anticodons, why do only tRNAs with three-base-anticodons exist in today's translation process? How is it prevented that other tRNAs exist and are used in the translation process?
Why don't we use a set of three- and five-base-codons together? Have tRNAs with five-base-anticodons been sorted out by natural selection because of a higher failure rate? But having longer codons/anticodons could also mean that failures are easier recognized, doesn't it?
Are there cases where tRNAs with longer anticodons lead to wrong translations? In the article tRNA hopping is mentioned. Is tRNA hopping the same as having a tRNA with a longer anticodon or is it more like having a normal tRNA and ignoring some nucleotides?
I appreciate any thoughts, explanations or links to more literature on the topic.
Thanks in advance.
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I don't think there is any "reason" why most things in biology turned out the way they did. Surely there were not dozens of different origins of cellular life on earth using various genetic codes etc. and the one with a 3-base codon code won out. All evidence points to life on earth all sharing a single common ancestor. In contrast, there are some good "reasons" why computer code begins with binary data, divided into 8-bit bytes, etc... Likewise, most human spoken languages have vowels and consonants, but there are some languages with click sounds and other alternatives. There are several ways of creating written symbols to represent the spoken languages.
Given four DNA (and RNA) bases, and 20 amino acids, the three letter code makes a lot of sense. But there is no real reason life could not have used 200 or 600 different amino acids instead of 20, and no reason to have 4 bases instead of 2 or 20. If there were 6 or 10 bases and 200 amino acids a 5-base codon system may have been worked out.
Within the coding scheme that did evolve on earth, there are a lot of interesting questions. For example the 64 codons are not equally divided with each of the 20 amino acid having 3 codons, plus 4 extras for stop codons. Why do we have 6 codons for some amino acids and only 1 or two for some others? Some bacteria have close to 80% G+C genomes (20% A+T) and others are very A+T rich and C+C poor, which presents challenges for maintaining protein sequence conservation. In evolutionary studies, the third codon position is often considered to be "silent" and have very low selection pressure, but in many cases there is indeed quite a bit of selection on the "silent" sites.
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rRNA and genes that encode RNA proteins, although highly conserved, are used for phylogenetic analysis of distant species. Why is tRNA not recommended for determining phylogenetic relationships?
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We commonly use tRNAs in mitochondrial phylogenomic studies. Of course, not individually, but as a part of the entire concatenated dataset. Especially in taxa where the architecture is highly conserved, as it is really easy to "pair" them.
Things get a bit complicated when there are duplications and/or missing genes, but a part of the tRNA dataset can be used even in such cases.
In some cases they can improve the phylogenetic resolution. If I remember correctly, Cameron found it to be the case in insects.
Here are examples of papers where all mitochondrial RNA genes were used:
And this is an example of a situation where there are duplicated missing genes:
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Do bacteria with fast doubling times evolve faster than bacteria with relatively slower doubling times?
Has this been measured? I would suspect that there is a correlation, but I wonder if necessarily so.
I have just stumbled upon a case in which two strains of closely related bacteria (16S = 98.4% sequence identity), which are well known to grow tremendously slow (in the lab) appear as if they are displaying substantially higher rates of evolution than expected (e.g. other ortholog proteins in these strains are a lot more divergent than in other strains with similar 16S level of identity).
The other possibility I can think off is that they are actually evolving tremendously slowly, in such a way that the level of sequence identity between their 16S belies their deep divergence, allowing more time for other proteins to diverge further given their extremely slow doubling times (e.g. weeks rather than hours).
Any good references on this topic? What do you think? Thanks.
All the best,
Tanai
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It's quite likely that organisms with higher doubling time will have greater sequence homology and evolutionary conserved sequences. The chances of introduction of the mutations are more in genome when doubling time is small. Your observations are really valid!
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Please make your valuable comments regarding the above discussion.
Society science engineering biology genetics artificial intelligence life science evolution sociology psychology behavior medicine biotechnology warfare population earth civilization
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The world has currently reached the peak of its development in all fields. However, there is a growing fear of the risks that war between the super powers may erupt if so extreme or irrational people assume power and take decisions that may lead to the destruction of human civilization. Despite all this fear, we look forward to the prevailing of stability and peace all over the world.
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Will the concept of science-fiction genetic experiments to recreate the long-extinct dinosaur species used in the plot of the film "Jurassic Park" ever be possible?
The plot of the film "Jurassic Park" directed by Steven Spielberg is based on a simple, but currently unrealistic concept of laboratory testing of the reproduction of long-extinct dinosaur species.
The collected genetic material of dinosaurs from the blood of a mosquito sunken for millions of years in amber is the main material on the basis of which extinct dinosaur species are recreated.
The genetic material obtained in this way introduced into the germ cell of modern reptiles in the film gives the possibility of reproduction of extinct reptile species.
This idea is based on modern research and genetic experiments carried out in laboratories, whose aim is to create, for example, new crop varieties or produce drugs for specific diseases.
However, the reproduction of long-extinct species such as dinosaurs is still not possible because the genetic material undergoes deep fragmentation over millions of years.
The genetic chain of chromosomes breaks down into very short fragments. So short that there is no information on how to assemble them into whole chromosomes and the lack of enzymes that would be able to fragment these fragmented dinosaur DNA pieces into whole chromosomes.
But the technology of genetic research is developing. The whole genomes of various species of animals, plants and other life forms are studied. The knowledge base of genotypes and related species is successively growing in the Big Data resources created for this purpose.
Therefore, the question arises: Will the fantastic research concept applied in the plot of the film "Jurassic Park" ever be possible? Will it be possible to recreate long-extinct animal and plant species with the help of subsequent generations of research in the field of genetics in the future?
Will it be possible to create a real Jurassic Park in the future, within which dinosaurs will run among the vegetation composed, among others, of flowering and woody ferns, horsetail and ferns, or the restoration of the ecosystem from millions of years ago?
Or maybe a man should not even try this type of other than present ecosystems to play?
Is this also a matter of ethics? Is it not threatening modern ecosystems to restore ecosystems over millions of years, ie consisting of many long-extinct species of plants and animals?
Please, answer, comments. I invite you to the discussion.
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Hi there!
That's actually a very nice question!
First of all, I am not the best person to answer this, but I would suggest you the book "How to clone a Mammoth" by Beth Saphiro. This is a great reading that answers a lot of your questions!
The main issue with the idea of using fossils to obtain genetic material is that "fossils" usually have organic parts substituted by minerals (processes called 'permineralization' and 'recrystallization'). Therefore you cannot extract DNA from these fossils because everything was substituted by minerals. It is also true that recent studies seem to have found traces of soft tissues in extremely well preserved fossils, but this is still extremely far from allowing to obtain the genome of dinosaurs (if you are interested in this, check the works of Mary Higby Schweitzer --> https://en.wikipedia.org/wiki/Mary_Higby_Schweitzer#cite_note-18).
However, the movie played it quite smartly on this topic, suggesting that the blood was actually encased in the amber, therefore not really fossilized. Nonetheless, in the book cited above, they explain how they tried to obtain even the DNA of insects from amber (I seem to recall they were working on bees?) but always failed. Again, a team of researchers managed to found "emoglobin-derived porphyrins" in a mosquito encased in amber ( ), but this is still far from Jurassic Park, still.
What you can extract DNA from, is frozen material. That's how they managed to obtain the full genome of the wholly mammoth (in 2015) from frozen specimens unearthed from the ice of Siberia. And yes, there is already who is thinking of using it to inseminate elephants in order to be able to have their "Pleistocene Park" (I now it's in Siberia but I do not remember additional information, again, check the book!).
This obviously generated a lot of discussion on the idea of how ethical it would be to use elephants to breed mammoth.
Sorry for the long answer, but I truly find this topic extremely exciting!
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(THE ANSWER: I asked the question and now, on June 30, 2019, it is answered to my satisfaction by the knowledgeable people who contribute here and especially by Aleš Kralj, Jun 28, 2019 , who asserted that evolution is fundamentally unpredictable. The Theory of Evolution as it is used here refers to “macroevolution.” A theory is expected to both explain and predict; the theory of evolution only explains. The theory of evolution is not a theory and it is suggested calling it a working hypothesis: the working hypothesis of evolution.
With the question answered to my satisfaction, I am now leaving the discussion. You may wish to continue.)
Science is based on observations. Empirical relationships between observations are called laws. For example, a graph of the position of an object vs. time is an expression of a law.
Theory is based on things you cannot measure; for example, one cannot measure momentum or energy, but only calculate them. If the imaginary things can be used to predict laws or observations you have a theory. If your theory cannot predict something that is observed …. or predicts something that is not observed . . . forget-about-it..... the theory not the observation! If your theory can not predict anything, the theory cannot be tested . . .why did you bother in the first place? Theories come and go. Scientists hide theories that do not work. ... What does oxygen mean? . . . is oxygen really needed to make an acid? That theory has come and gone. Where did phlogiston go?
Do you believe in the Theory of Evolution? What can you predict right now from the Theory of Evolution? What new species will be discovered?
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@ Md Zafar Alam Bhuiyan
@Hamid Gadouri
@ Everyone and Anyone
Here I would hope for a discussion based only on “natural” philosophy. Not a discussion that in anyway involves theology, or any other school of knowledge.
Basically is there a “Theory of Evolution” or only empiricism?
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Does anybody know a lab in Australia or New Zealand who carries out in situ hybridization, FISH, GISH with polyploid plants?
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I think in Wellington Regional Genetics Laboratory, CCDHB New Zealand
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Adaptations correspond to physiological ranges (reaction norms, somatic adaptations). According to the theory of facilitated variation, such dynamic physiological restorations of the phenotype in response of variable environmental conditions are the outcome of genetic constraints (e.g. plasticity and robustness of developmental pathways). Therefore, when somatic adaptation occurs, exposing the phenotype to different selective conditions, physiological ranges can be "easily" shifted (i.e. their evolutionary shift is facilitated) by mutation, or genetic reassortments from the existing variability in the population (Baldwinian evolution). In other words, one of the key characteristics of adaptations would be their evolvability, or to say this with the words of Gould and Vrba (1982), "cooptability for fitness". Evolvability can thus be strongly conserved at the level of core molecular processes. Adaptations would then be selected to be both physiologically adaptable, i.e. to function in "a range of ways" in response to changed conditions ("dynamic restoration" or somatic adaptation), and to be evolvable. In other words, the "cooptability for fitness" would be under selection.
In my view, this idea implies that all adaptations at the organismal level should be partly selected to be "preaptations" (sensu Gould and Vrba 1982, Paleobiology), i.e. structures that retain the potential to enhance fitness (adaptive function) in variable conditions. Gould (2002, the structure of evolutionary theory) suggested that this selection should act at higher hierarchical level (species selection). However, the fact that adaptations that are selected in specific conditions are also selected to be physiologically modifiable, or to function in “a range of ways”, would make them likely to have fitness-increasing effects (aptations) that are not those they were selected for during their historical genesis, i.e. to become exaptations. Could this be a bet-hedging strategy also selected at organismal level?
That is, exaptations would also have a non-random origin (contra Gould & Vrba 1982), while this does not rule out the possibility of non-aptations as a possible source of exaptations. In this scenario, exaptations from nonaptations (spandrels s.s.) would be less frequent than exaptations from previous adaptations. Note also that the measurable adaptations are a subset of the extant ones, due to the overall scarcity of available historical data.
I will often modify my comments.
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Gianluca,
Thanks for the feedback. My comments below:
1. Pluralism in the MS just means acknowledging measured contributions of the main evolutionary forces: selection, drift, gene flow and mutation.
But there is no mention of mechanisms for the origin of mutation and their spread through populations without selection, drift, or gene flow.
2. Either a structure has an effect on the fitness of the organism, then being either maladaptive or adaptive, or it hasn't, thus being non(ad)aptive. In the latter case, it just does not have any function (neutral).
If you wish to define function that way.
3. Exaptations (shifts of function from either an adaptation or a nonaptation) are real stuff, not dreams.
Not so sure about that.
4. Gene duplications, pseudogenes and genetic shifts of functions are the results of historical events, not dreams.
Sure, but the dreaming comes in when people invoke ‘selective pressures’ or other such without any evidence.
5. Bird feathers are …the end result of a historical process made of contingencies and interactions between biological systems and environments:
true of anything (and anyway, a feature is really two feathers in one)
6. Gould ….. aimed to contrast adaptationism by reinforcing structuralist themes, which seems quite the opposite of what you're saying.
OK. Although I think he just further muddied the waters of Darwinian evolution. But that is just a personal opinion.
Cheers, John
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Dear Thanos,
Thank you very much for the congress activity, impressive development and my invitation.
I finalizing my paper to the congress issue: Yuri Kartavtsev. "The evolutionary-and-genetic paradigm development: Neo-Darwinism vs. molecular evolution fit”. It is an assignment, short as possible. However, it in demanded format is 8 pages without References and 3 figures. Total volume is 17 pages. Should invited speaker follow 5 pages standard or there is a relaxed limit in this case?
If you will accept such MS volume, I’ll proofread finally this short review and submit it soon after on the congress site as requested.
For an occasion I attach draft of the paper.
Cordially
Yuri
07.04.18
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Dear Michael,
Thanks a lot for nice news. Always there a time limit...
With warm regards
Yuri
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I've been told to make a selective pressure analysis of a hypervariable region of a viral genome. I have aligned my sequences, removed stop codons and I did some tests I've met in publications like Codon-based Z-Test of Selection in Mega 7, MEME, FEL, SLAC methods on Datamonkey server, and tried to mesure synonymous versus non-synonymous mutation ration with the SNAP program. But I have absolutely no idea what to do with all the obtained numbers and how to interpret the results. Can somebody simply explain the selective pressure analisys to me?
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for selection presure analysis you can read this book that might be helpful:
Molecular Evolution and Phylogenetics
Masatoshi Nei, Sudhir Kumar
Oxford University Press, 2000 - Medical - 333 pages
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I have once been asked about the methods for studying ancient DNA, and I am rather naive for knowing too little about studies on Ancient DNA. I am hoping to receive more information about sequencing Ancient DNA (Again other than mtDNA and Chloroplast DNA) for studying ancient samples. In particular Human samples.
(Assume we're gonna study them through pedigree diagram, are there any more specific methods for studying the Ancient DNA samples: Such as Haplotypes)
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Thanks for replying and I'm really sorry about the late reply!
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I am trying to identify and elucidate the function of a gene (let say X) in Brassica napus that control flower development in Arabidopsis. Using the sequence of Arabidopsis X, I conducted a phlyogenetic analysis based on protein sequence data. Phylogenetic analysis revealed that there is a single X in Brassica napus genome. After phylogenetic analysis, I isolated the cDNA sequence of X from Brassica napus by RT-PCR and then sequenced 20 randomly selected clone. When the coding sequences of these clones were compared, they revealed two unique sequence which were highly homologous except single nucleotide variation in some position of the coding sequence. When the two unique sequences of Brassica napus X were compared to its progenitor (Brassica rapa and Brassica oleracea X), I found that both of them are highly similar to the sequence of their progenitor X both at nucleotide and amino acid sequence level. The identity between Brassica napus X and Brassica rapa X were 98% (nucleotide level) and 93% (amino acid sequence). The similarity between Brassica napus X and Brassica oleracea X were 100% both at nuclotide and amino acid sequence level. My question is
1. What will be the possible explanation of presence of a single Arabidopsis X in Brassica npaus genome given the fact that Brassica napus is a tetraploid crop?
2. Are the two unique sequences of X found Brassica napus genome represent two different allele or two different genes?
3. I would also welcome any other explanation on this result.
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To really determine how many copies you have and if they are homologs, you will have to take one of two approaches.
1. Old school - Southern blot with Gene X as your probe. This will tell you how many copies of Gene X there are in the B. napus genome.
2. Bioinformatics - IF there is a high-quality genome sequence, you can look for the copies of Gene X as a prediction. Then, clone and sequence these regions from B. napus
Or combine both approaches. Good luck!
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I am trying to understand the molecular evolution of a gene family that composes of only two members (let's say X1 and X2). I have looked into the genomic neighborhoods of X1 and X2 in various species using Genomic Alignments in Ensembl and constructed a genomic neighborhood illustration of other genes flanking X1 and X2. However, my Blast searches didn't detect X2 in some fish species. When I looked into the genomic neighborhood of X2 in these fish species, I saw another X1 like sequence instead. I am thinking that maybe X2 was converted into X1 after the duplication event, but how can I prove that? My assumption may be stupid because I am not very experienced with genetics and concepts of evolutionary biology, but any help would be appreciated.
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Your assumption is reasonable. Another possibility is simply that one of the genes has been lost. Some fish species have had 2 rounds of genome duplications and afterwards lost some of the duplicated genes. Doing a synteny analysis, as Abhishek Kumar also recommended, would enable you to distinguish between gene loss and gene conversion.
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Since amino acids are degenerate in nature, the amount of variation will be higher than its respective nucleotide sequence so will it make sense to construct a phylogeny tree using the protein code?
Also, how different should the sequences be from each other to be marked as a different lineage or under a different internode?
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As usual in science, it depends on the aim of your study. If you are trying to unveil the taxonomy of an unknown species, I would suggest you to work with 16S for bacteria/archea and 18S for eukaryotes.
If you are studying protein coding genes, as mentioned by Dr. Kumar, it might be more reliable to work with aminoacidic sequences (or translated nucleotidic sequences), as they have a higher information content ( 5x fold).
About the threshold of similarity in clustering sequences, it mostly based on experience on the sequences you are working with. It depends if you are working with nucleotides or coding sequences, the expected divergence between sequences, the evolutionary rate (K) of that region and the evolutionary distance between organisms that harbour the sequences. On top of that, there is also the aim of the study. It is not the same comparing variations among closely related populations (high similarity is expected) than variants between sequences that belong to different orders.
I would suggest you to read the books and reviews of Dr. Felsenstein, to learn the best approaches to your problem, as well as the most common pitfalls.
Cheers.
JMC
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I need to reconstruct ancestral sequences in evolution of short peptides. Trying all possible models in FastML I obtained results that just didn't make much sense. In Mesquite there are currently no available likelihood models for protein evolution... 
For me this is a first experience in this field. You, who do have some experience with peptide (or more likely protein) evolution reconstructions, which programmme would you recommend to use?
Thanks
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Use MEGA7. The King of them all
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I have a SNP dataset that contains several cryptic species. I can separate the individuals into groups using STRUCTURE and PCA, but I'd like to give them a nice number so we can say that yes, they are actually substantially differentiated.
For my last project we were just using single sequences, so I could calculate plain old sequence divergence. Is there anything like this for SNP data?
Alternatively, I do have the sequences the SNPs were retrieved from. Is there a way to calculate divergence using many sequences?
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Hi Emma,
 If you want to calculate the genetic diversity of a single population, there are several estimates that you can use such as the allelic diversity, proportion of polymorphic loci, observed and expected heterozygosity, and gene diversity of Nei (pretty much the same that you use for sequence data). A very friendly software that estimates all these parameters is GenAlEx (http://biology-assets.anu.edu.au/GenAlEx/Welcome.html).
 Now, if you want to quantify the level of population subdivision, as Manolo suggested, F-statistics are more appropriate. PopGenome package is one of the best alternatives in this regard, although there are also other free available softwares such as Genepop (http://genepop.curtin.edu.au/) that are also pretty easy to use.
Cheers,
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I was wondering what people’s thoughts on this topic were: how many SNPs per RAD locus should be used for phylogenetic studies?
All agree this is important, with fewer SNPs per locus being considered more ‘robust’ or ‘true’, but I can’t find a real good study looking into it. With my own data the strongest supported trees were generated when I filtered out all loci with more than two 2 SNPs (range of 2-10). This does not make them more ‘real’, but does have an impact.
iPyRAD is interesting, as its default is to allow 20 SNPs per locus with a minimum read length of 35.
So what do people think? Are their good studies on this? How are you determining the best maximum number of SNPs per locus when filtering data for your studies?
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Hi James, in my experience I gain good results by using not more than 5 SNPs/Locus. It also depends on the level you are working on. It might be a difference if you are working on closely related or distantly related taxa. I am working within a tribe and subgenus, resepectively, with good results using only up to 5 SNPs. A setting which allows many SNPs will increase the likelihood of including loci that might be the result of paralog alignment/ sequencing errors/ gene families and so on - despite the filtering beforehand. This will lead to "noise" and contradicting results in the calculations leading to lower BS support. But again, this might be different if working within a species on populational level. You can also try using the unlinked SNP files (one SNP per locus) to test the differences in your results. Good luck!
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Is there any difference in them molecular level? Even though it is produced from different combinations of genetic code but produces the same amino acid, is it acceptable between eukaryotes and prokaryotes ? 
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If you are asking if there is any difference between glycine coded by GGA and glycine coded by GGC. The answer is 'no'.
In certain different systems, the codon table varies.  eg AGA is arginine generally, but a stop codon in the mitochondrial genome.
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For divergence time estimation
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Hi,
There is no generally accepted substitution rate. You can calibrate a sutitabele substitution rate for your target marker using geological events or fossil records in BEAST software. Hope this helps.
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I need to reconstruct the nodal position of some fossil taxa on molecular tree with morphological data about extant species. My data have quite complex structure; categorical are multistate (sometimes contain ambiguities) moreover other are continuous. I want to ask if EPA implemented in RAxML:
1) Will deal with ambiguous states? In other sofrware I have often had a problem when trait encoded as e.g. "1/2" was reconstructed as a separate state.
2) Will deal with continuous data? Do they require categorization? I will have to that arbitrally, because there are no natural categories in some of traits (basing on analyses of their distribution). 
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Thank You for Your answers! I have dug through litearture and currently there is practically no way to deal with compund morphological data. Albeit polymorphisms can be simply encoded as dummy variables, there are completely different models for continuous and categorical data. I have decided to use Gower's dissimilarity index - just a simple phenetic but there is no other option if You are unwilling to categorize Your data. 
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Hello,
I want to sequence rRNA16S of bacteria  by means of next generation sequencing technic using Illumina MiSeq  technology. In articles i see that they choose hypervariable region V4 or V6 ..knowing that they are 9 hypervariables regions in rRNA16S.I want to know on which criterion is based the choice of the hypervariable region of bacterial rRNA 16S ?
Thank you
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This Nature review may provide some insight as to which variable region to choose.
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I could find median and 95%hpd in figtree.
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Hi Ashwini,
Typically, you would first need to define your ingroup in BEAUti's Taxon Sets tab. After running your analyses, as Ana mentions, you would upload your logfile (.log) in Tracer to look at different statistics of the time-to-the-most-recent-common-ancestor (TMRCA) of your ingroup.
In the event that you did not specify a taxon group in BEAUti, and you are unwilling to repeat your analysis, another option is to use R:
For instance, you could use the ape package (read.nexus function) to upload the whole posterior distribution of trees (your .trees file). After discarding the burnin, you could loop-search and extract the node age of your ingroup. Once you have the node ages in a vector, you can do whatever kind of statistics on it (e.g. using mean() and std() functions). This approach requires a bit of knowledge on how to deal with tree data in R, and R in general. Have a look at the attached link if you are interested.
Cheers,
Santiago
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Dear all,
I'm having trouble on producing binary trees in PAUP*. It seems I can only do it by Trees/ Generate Trees. But when I do this it seems that I cannot 'use' the already produced trees but that I have to re-generate them (or am I getting all these wrong?). Any thoughts?
Many thanks,
Toni
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I should add that when you use generate trees, you are producing a set of random trees that are not based on the data at all. that s whether are binary. This method does not find a best or optimal trees from your data. The command is used to help you look at the distribution of tree scores to make sure that only one or a few trees have support from the data. Let me know if you do not understand this.
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I will be on the field soon (in Madagascar), and will need to collect RNA to get a transcriptome. Of course, I won't have access to liquid nitrogen. Thanks for your answers and advises
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you can use RNAlater Solution
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How does a tissue sample need to be preserved in order to do whole-genome sequencing? Interested in preserving opportunistically acquired samples for future analyses but I'm not familiar with the needs for preservation.
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One easy, reliable and economic method is to preserve the tissues for DNA with higher integrity and further extraction is in ethanol 70% (liquid exceeding the volume of the tissue 2X) in a normal freezer (-20ºC). 
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i am planning to use Primer v6+PERMANOVA to analyze metagenomic data, any one have this experience? what is the difference between analyzing metagenomic data and morphological data using this software?
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Keith McGuinness from CDU has a Youtube playlist with a series of videos on how to analyse amplicon sequencing data and it's nicely explained (see link)
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how to calculate dn/ds ratio for list of mrna and lncrna from mouse data against human?
I have used biomart from ensemble and get list of orthologous against my data and dn/ds ratio? how I can get accumulative result or make accumulative this result from this data like
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Dear Azhar sahb,
The PAML package here are different tools inside. out of them the yn00 software, that compute dN/dS in a pair-wise manner between two sequences:
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We are having demo projects on integrative systematics on bees and wasps in the lab. To gather enough data, we would like to try following technologies of next generation sequencing - 
RADSeq, Restriction-site associated DNA sequencing;
AHE, Anchored Hybrid Enrichment;
GBS, Genotyping-by-Sequencing;
UCE, Ultraconserved Elements.
Any comments or suggestions? 
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Dear all,
We are doing RADseq on Trichogramma unique individuals, these "beasts" are ca 0.3 mm long !!!. To do so we use Qiagen individual extraction followed by WGA and it works very well. We tested the reliability of WGA and we did not detect any problems. To do so we let a male (M1) and a female (F1) tricho mating. We obtained multiple female descents (F2s) that we isolated (no mating) and let them produced males (M3s). We pooled all these males (n=ca 2000, in a kind of natural genome amplification) and compare the results with WGA of  M1 and F1. However, for some reasons we had poor results using WGA for UCE capture, in average we lost one third of the loci captured on WGA individuals.
Best
JY
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We have detected a rare mutated sequence and I am trying to find out if the sequence is absolutely unique or it has already been observed. I blasted the sequence and did not find any matches in the free public databases like NCBI and UniProt. The sequence resulted from in-frame deletion of several dozens of the base pairs.
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Thank you, I will definitely try that.
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Hello, I would like to confirm that it is possible that ITS2 ribosomal DNA marker may indicate introgression and hybridization through inferred trees with this gene?
The tree recovered with this marker showed a polytomy with short branches between two sibling species, can I conclude that it is evidence of introgression only with this marker, in case the ITS2?
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Hi José. Usually polytomies indicate lack of resolution in your data.  A hypothesis of hybrid origin (introgression) cannot be rulled out, but the short terminal branches may also indicate weak support for a hypothesis of introgression. Why do you think that your particular tree indicate a hybrid origin of the ITS2? With a single marker it is easy to manually verify the origin of each mutation. For each polytomy select three specimens representing the suspected parent (monoclonal) populations, and the suspected hybrid, and visually compare their aligned sequences. Make a table classifying the various types of nucleotide similarities: positions with nucleotides unique to a each specimen, and positions with pairs of nuceotides shared by each of the three possible pairs of sequences. For ambiguous base calls (presumed polymorphic positions) consider the individual nucleotides comprising the ambigous call (but check the chromatograms to make sure that the polymorphisms are real and not the result of poor quality chromatograms).  If the frequency of pairs of shared nucleotides are large and close to 50%, 50%, 0%, the hypothesis of hybrid origin is preferred; otherwise alternative hypotheses, such as lack of ancestral resolution or stable polymorphism may be preferred.
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Hello. I am deeply puzzled by the breaking-news study by Fennessy et al. (2016) "Multi-locus Analyses Reveal Four Giraffe Species Instead of One" (dx.doi.org/10.1016/j.cub.2016.07.036). The nuclear analysis seems to be solely based on 7 introns sequences, and contradicts a previous study, by Brown et al. (2007) "Extensive population genetic structure in the giraffe" (dx.doi.org/10.1186/1741-7007-5-57), which included over 3 times more individuals for 14 microsatellites loci, with samples from contact zones, and found up to eleven subpopulations clearly differentiated from a nuclear point of view (with possible hybrids)... without suggesting any taxonomic emendation though! As results of both studies are not clearly confronted in the 2016 paper (I barely found a terse "the statistical support is not clear" when mentioning the 2007 results), could any one explain me where are the new insights brought to the Giraffe's case by the latter? And who to follow?
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... And you will find some people loooking and looooooooooooking to all genes of their dataset until they find THAT marker (in the middle of a neutral global result) that will show that HIS/HER SPECIES is statistically "real" (for example, frequencies of an allele coding for color under a strongly selective environment for that color). And the worse part: they get published in more-than-good journals. Now with RADs, for example, you could "find" a species almost everywhere, if you look enough.
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I'm a bit suspicious to that tree, to me it looks odd to have A0 being separated from Aalba with such a relatively long branch and still not have Aalba monophyletic. What methods is the tree estimated with and is it correctly rooted? I would guess you may get a different result if you reestimated the phylogeny with a different method and if needed additional outgroups. But as already said Aalba is paraphyletic and A0 is monophyletic in your tree
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For examples in molecular cloning of a gene fragment
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As of December of 2015, the American College of Pathology and the American College of Medical Genetics published guidelines regarding this matter. The terms mutation or polymorphism, should not be used. Now the term is "variation" that can be either, pathogenic, benign, or possible pathogenic or possible benign. In order to know in which category a variation should be classified, needs to be analyzed in different databases in order to establish if it has been found  associated to the disease. Databases such as snpdatabase, mutation database, EXaC database, 1000 genomes etc, should be searched. Also different prediction softwares such as polyphen-2, mutation tester,  provean and PHDsnp should be used.
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I want to know whether speciation rate of any particular insect family is larger than others. How can I do that? What are the simplest and suitable bioinformatic tools to address the question.
Thanks 
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pretty much as above, probably want to check out what is available in the lit also!
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I'm trying to differentiate between species in the samples I have and would like to sequence the first 700-800bp on the 16s rRNA sequence first (using sanger sequencing and the primers 27F plus some universal reverse primer), then send in the last 800 bp for sequencing (using 1492R and some universal forward primer) if the first try doesn't get me down to the species level. It's been difficult, however, to find internal primers that are universal or have a link to a paper using them successfully. Anyone know of any?
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Thanks David, that is an extremely helpful response.  I was able to open the alignment file in a MASE and determine based on a similarity threshold what areas are most variable. This should help greatly in designing any primers for metagenomic studies in the future.
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Hello dear colleges!
Well, I have this issue that are driving me crazy. Im trying to amplify COI Folmer Tailed HCO/LCO primers for Barcoding, but I have this faint bands (Attached file). I already try changing the Thermal Cycler (in fact I already used three different thermal cyclers), PCR conditions, PCR program, Quantity of DNA, Quantity of primers, MgCl, temp grad, and more... The DNA is working because I already amplified 12S and 16S
Other thing is that I don't know why my ladder is faint too, I already bought a new one and is the same, I used another power supply and different % of agarose and nothing change, we are using Et-Br, because we're out of gelred stock.
Any suggestion?? I'm little stressed about this :( 
Thank you everybody!
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I think there might be a problem with running buffer. Use freshly prepared 1X TAE ( use Tris Base not Tris buffer while preparing buffer). 
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I want to identify two different individuals of parasitic plants growing in one host from each other. Thing that I know, to do DNA barcoding for species identification we can use mitochondrial cytochrome oxidase I gene (COI). Is there a DNA identification protocol for individual level? Are we using specific genes as markers?
Please let me know if there any reference for it.
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Dear Arthur Burzynski,
Thank you very much for the orientation and contribution. I find this kind of discussions helpful for my science, although there may some occasional misunderstandings occur.
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Dear colleagues,
I want to sequence some genes from different mammals species. However, the major part of them has no reference sequence. What is the best strategy to do it? Is there any free available software that I can use?
Thanks in advance for your help!
Best,
David
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I have had to do this myself and it's a pain. The best thing you can do is search databases (NCBI, Ensembl) for sequences of the closest relatives you can find. Put them all together in FASTA format in a .txt document and do a multiple alignment using Clustal Omega. It's best to do these alignments using mRNA/cDNA transcripts or complete coding sequences (cds) because introns will not align well. However, you will need to design primers within exons (not across junctions) if you want to sequence DNA. I do this by using Ensembl to figure out where exons are and using the longest exons as the input to primer design programs (like Primer3). Of course, if you can't get a section long enough for your purposes you will need to use Ensembl to look at how far apart the exons are and choose a pair that give good amplicon length, then design primers in each exon individually. I'd recommend excluding sections of the exon that are not well conserved according to your Clustal alignment.
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Hello everyone
I encountered this issue while extracting DNA from mollusc tissues with a salt-precipitation protocol. For some samples, the pigments contained in the tissues (which were originally black, grey, red) remained in the fraction containing the nucleic acids during the whole procedure. The pellets of DNA resulted coloured at every step, and the final elution maintained the same color. In my case the final elutions were of different shades of yellow, coffee-like brown, and red (always very bright), but I heard of similar cases with other organisms where the final elution was green.
I don't know the nature of these pigments in my animals (i.e., what kind of molecules they are), and all I can think about is that they may at least have the same polarity as DNA since they move in the same way during the extraction procedure.
Do you know of some possible negative influence of these pigments in downstream analyses? If yes, is there a way to separate them from the extracted DNA? For now, I did not see clear effects on quantification with a spectrophotometer and in PCR (colored samples behaved similarly to clean ones), but I don't know if in qPCR the pigments may alter the fluorescence signal.
Have you ever encountered similar issues? If yes, I will welcome every comment you can give me! Thank you!
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This problem is frequent for people who work in plant sciences (molecular biology, biochemistry). Most plant pigments are phenolic compounds and we get rid of them with PVP (soluble or insoluble) during lysis steps. These are polymers which chelate phenolic compounds before they make links with nucleic acid (and reduce enzymatic activities). I'm not familiar with animal tissues but I just read that some of them contain pigments with phenolic radicals (melanin). Moreover, hemoglobin-derivated pigments can be fixed on Polyvinylpyrrolidone. And these pigments are also known to reduce qPCR efficiency.
I think the use of insoluble Polyvinylpyrrolidone is the easiest (and cheapest) way to chelate phenolic or hemic pigments, you can use it during the lysis step of commercial extraction kits : the chelated pigments sediment with cellular by-products and solubilized nucleic acids can be processed during further column-based or beads-based purification steps.
Jai Ghosh points another way to improve nucleic acid purification : it seems that invertebrate contains proteic pigments and in theses cases, a PVP-based step would be inefficient.
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Cost is more critical than time ;-)
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So I always used the old school phenol:chloroform:isoamyl alcohol (25:24:1) technique. It's from Current protocols but I've attempted to attach below my write up of the protocol from grad school. You would start around step 15. Make sure whatever protocol you use has the isoamyl alcohol as this is a substantial improvement over even older protocols. As you're on buccal brushes, most people I know soak the brush in TE buffer and then you can proceed as described but other protocols I've done, for the buccal swab (I wasn't actually using a buccal swab) would mean add your TE, then add phenol:chloroform:iso to the TE with the brush still in and mix, vortex, or soak. Then proceed as described. As in this written protocol, you'll definitely get higher yields of DNA if you precipitate over night at -80. Glycogen will reduce the odds of you losing your pellet. And I've done this same protocol on the 200 uL scale which was probably more consistent with what your DNA yields are going to be, but you have the problem of the large buccal brush that you need to sufficiently wash. Note in the below protocol, it's just the ratios of reagents to the starting amount of buffer that the DNA is contained that matters. If you want to soak your brush in 1 mL you can adjust everything accordingly. The least amount of DNA I've purified reliably and consistently with this technique is just below the nanogram levels, so 100s of pg. I've simply not tried even lower, but it does take a little bit of finesse. Hope this helps.
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Is there any software (desktop or online) that allows you to create customizable visualizations of the most current phylogenetic relationships between taxons? Let's say that I would like to have the illustration of phylogenetic relationship within extant mammals and extant birds and I would like to include different detail, like at the level of orders or at the level of families, or detailed relationship in selected order only. Then maybe change the layout little bit and export it into word processing software or save as vector graphics for further editing. Is there something like that? Something easy that students can use for the assignments and theses, etc. I think it would be handy if this would be possible in Tree of Life but I was not able to find any feature like this there. Do you have any ideas? Thank you!
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Dear all, it has been a while since I posed my question and I apologize for being late with my own response. However, I finally got to look at your valuable suggestions. With your help, I learned few new things and got perfect answer for my question and even beyond that question.
I would like to summarize a few points:
Livio suggested www.timetree.org and I was able to find there pretty nice dendrograms for some of the taxons and very nice poster of tree of life. However, I think it is not possible to adjust the dendrograms in any way. Still it is good for the purpose I originally asked for and I think students could use it to illustrate phylogeny of their group very nicely.
Brian, Agostino, David and Eduardo suggested different software that can be used to visualize the trees. ETE Toolkit looks very very powerful but I could not find Windows version (just MacOS and Linux). In future, I hope to use OneZoom as it is indeed customizable (allows uploading own trees and metadata) and very attractive for public presentations. Eventually, I used FigTree which is powerful and very easy to use. I also found Interactive Tree of Life http://itol.embl.de/ which looks also nice. However, I needed to get the tree in text format to upload in these programs first...
Adam suggested Open Tree of Life. I was not able to find how to export the nice dendrogram pictures directly at the site but I found it is powerful to get the data for other programs. It took me a while to learn how to work with API (eventually, I found a good instructions here: https://blog.opentreeoflife.org/2014/10/07/tree-for-all-hackathon-series-introduction/ and used DHC Chrome extension to run API orders) and now I am able to extract the current dendrogram even for the small set of species I am only interested in and which are scattered across all vertebrates and then make a nice dendrogram in FigTree. Great for visualization purposes!
Thank you all! Best, Pavel
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Which one is suitable for phylogenetics study: mitochondria genomics or chloroplast genomics? To construct trees.
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Hi Bihua Chen,
I think it depends on which group you are working on. For plants, the evolutionary rate of the mitochondria is very low, then the chloroplast genome might be a better choice. In addition, I agree with Manolo Perez, sample nuclear genome will avoid inappropriate interpretation based only on the cytoplasmic genomes. Hope this helps.
Xiangqin
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in case of equidae family
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When you remake the tree with only the sequences used in the original source you are referencing, do you get the same result? Are you using the same model and parameters as used to generate the original reference? Assuming replicating the original gives the same answer, then the changes come from adding more data, which is unlikely to actually be a problem. As long as the extra data is not bogus, the addition of data often changes our results and interpretations somewhat.
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I have reconstructed the phylogenetic relationship of a kind of mammal using different mtDNA and generated different results. Specially, four consistent clades were resolved by different mtDNA, but the phylogenetic relationship among the four clades verified among cytb, Dloop and ND1. As you know, there are mountainous studies suggested that the phylogenetic trees generated from different mtDNA are usually same as the mitochondrial genome inherited as a single unit. Numts and insufficient sampling are not likely the case in our studies and related studies are rare.I am kind of stuck now and failed to find similar evidence, therefore I greatly appreciate your valuable knowledge and suggestions!
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You could run your tree and alignment through PhyDesign (web app) to better understand what the rates of evolution of markers imply about their Informativeness at different depths of your tree.
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References:
*Kalen et al. 'Age-related changes in the lipid composition...' Lipids 1989; 24: 579-85.
*Alehagen and Aaseth.'Selenium and Q10 interrelationship...' J Trace Elem Med Biol 2015; 31:157-62. 
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Together with Urban Alehagen, we used a combination of selenium and Q10, but perhaps only the selenium component was responsible for the preventive effect?
 Alehagen, U., & Aaseth, J. (2015). Selenium and coenzyme Q10 interrelationship in cardiovascular diseases–A clinician's point of view. Journal of Trace Elements in Medicine and Biology, 31, 157-162.
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Dear colleagues,
My colleague and I cloned a gene from sword bean which is mainly expressed in flower. After I did blast using the protein's amino acid sequence, I found there are just 250 homologues in the database (Uniprot), and only exist in angiosperms including the basal species  Amborella trichopoda.
Then, I used the amino acids sequence of these proteins built the
phylogeny tree as attached, and placed Amborella trichopoda at root.The tree (NJ) looks a little strange,  the monocots mixed with dicots. In fact, when using NA sequences to generate the tree, the tree shapes are similar. 
My questions are :
1. Because no homologues could be found in other than angiosperms.
Can I say this protein appear very later along with evolution of angiosperm? 
2. Can I say this protein evolves slowly or conservative, the 250 proteins from angiosperms range from 260-300 AAs and few subtypes of this protein had been reported ?  
3. Convergent evolution? Monocots and dicots mixed in the tree.
 
regards
 Zha
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Zha,
It is hard to judge unknown gene evolution, but using common cense, I would say/ask: 
1. The fact that no homologues have been reported  in other plants than angiosperms does not mean: 1) that the protein/gene is absent in other plants - it has not been reported (but still, can be with low similarity)  so far in the studied plants out of angiosperms lineage. 2) It cannot be said that this protein appear very late along with evolution of angiosperm - it  must be present in the last common ancestor of angiosperms, and it is very old event, and it could be present in other plants of that age.
2. Is the protein-coding gene present in a single/few copy and has no paralogs?  Angiosperms have origin about 247 million years ago - plenty of time for evolution of many genes. Your protein must be compared to other to make conclusions about evolution rate.
3. 'Monocots and dicots mixed in the tree' - again the question about the gene copy number. If there are some paralogs - it is normal.
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Is it as simple as: genes are only transferred from one taxa to the other and not the other way round?
Thank you
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Hey Rafaela! how are you... just saw this one...
basically, imagine two species, Blue and Red, forming a hybrid, called Purple.
Imagine somehow these purple hybrids are not able to form a new independent biological unit. So their individuals can only reproduce with individuals of the parental species. But not both! imagine that the Red males are not able to fertilise the Purple females, and the Red females die everytime they copulate with Purple males. This way, purple animals can only interbreed with Blue ones, by doing so, since they are hybrids, they bring portions of the Red genome into the Blue one. In the long run, because the Purple hybrids are not very fit, you will have populations of Blue that mostly resemble the Blue species, but which contain "genomic traces" of the Red species, introgressed ASYMMETRICALLY at the time of the backcrossing of the Purples.
the most common case would be when you detect a clearly "Red" mitochondrial haplotype, in an otherwise "blue" looking animal: that is a signature of this asymmetrical introgressive event, because even after many generations of Blue interbreeding, with the Blue nuclear genome being predominant, the mitochondrial lineage coming from that one "purple" female that mothered a backcross offspring is still there, being transmitted.
There are several cases, for example in blue mussels Mytilus spp. even us, humans, have a little bit of Neanderthal DNA... which I think is asymmetrical. well, the poor Neanderthal are extinct now, but some of their DNA lives within us!
I have simplified it a bit... but I hope it makes sense...
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* stress conditions usually results adaptive mutations
* adaptive mutations are realistic?
* in adaptive mutations alterations takes place at genomic level or phenotype level 
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Dear S.H. Ramaprasad, 
the term mutation refers to alterations in DNA sequence. And it depends on the nature of the mutation, if the mutation produce a change in amino acid, these radical changes are many times involved in changes in phenotype, by protein destabilisation/loss of function/neofunctionalization/subfunctionalization
When your organism acquire a single mutation, you can expect that these organism is a mutant, but you should also consider that bacteria usually form populations, and populations have genetic polymorphism, so you should be aware to not confuse mutant with a genetic variant.
Did you mean with "real bacterial mutants" that the mutation confers a new phenotype? 
There's a lot of literature on adaptive mutations in bacteria and the arose of mutants check for Richard Lenski and his 50.000 generations experiment; or those from Mario A. Fares in bacteria and yeast (with myself as co-author). 
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as with the field of phylogenetic inference, methods to resolve polytomies are being developed on species trees instead of intraspecific trees. my interests focus on intraspecific trees, usually using likelihood methods like the coalescent, and I have yet to find methods developed to test for hard polytomies in these sorts of trees. since species and intra-species population and genetic dynamics are quite different, I would expect the methods to be different as well.
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I have two DNA sequences (seqA and seqB) deriving respectively from two longer fragments (fullA and fullB) and a phylogenetic tree from which I can obtain the evolutionary distance from fullA and fullB (dist(fullA,fullB)).
I'm then interested in calculating the probability of the fragments seqA and seqB having less nucleotide dissimilarity than expected given the observed overall nucleotide dissimilarity between fullA and fullB and the evolutionary distance between them (dist(fullA,fullB) which is a proxy of the 'available time to generate dissimilarities').
Do you know of an implementation or model to calculate this probability? any reference to a published work addressing a similar question?
Thank you so much in advance.
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Dear Joseph,
Thank you for your answer!!! Indeed, Modolo et al. 2014 (http://10.1093/gbe/evu026) apply an approach similar to the one you described in your answer. They report that: the number of different nucleotides between the two sequences follow a Binomial(Ln,pn) distribution with Ln being the length of the pair of sequences of and pn the probability of having a nucleotide dissimilarity. A binomial test could then be applied to calculate  for each pair of sequences n the probability of having a number of different nucleotides lower than expected, or unilateral P-value.
Dear Pablo,
I will look at DnaSP to derive the expected nucleotide dissimilarities on the small sequence.
Thank you so much for the replies. 
Best,
Aminael
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I'm wondering e coli regulate some gene actions in primates in the so called mechanism. (Bacterial regulatory effects on primate genome )
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Hello,
That's a very good thought process.
We have recently published a hypothesis based on inter-specific miRNA transfer and possible expression.
The link to the same is: doi:10.3390/ncrna2020002
Happy reading.
Warm regards,
Venk
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I am interested on identifying signatures of balancing selection with ddRAD data. Any suggestion?
Thank you in adavnce
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Hi Francesca, I found this review useful to get the picture: "Detecting balancing selection in genomes: limits and prospects" http://onlinelibrary.wiley.com/doi/10.1111/mec.13226/full
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I've been doing molecular cloning for many years now and I've recently moved labs. In my new lab I'm having an issue with band splitting in my DNA gels, the lower bands in particular look letterbox shaped (see the smaller fragments of the DNA ladder in the first and last lanes for an example). These are all 1-2% agarose in TAE, stained with ethidium bromide. I've never seen issues like this in my previously lab and I can't think of an obvious difference in the setup here. I'm running short gels (7 cm) at 70-90 V.
Has anyone else seen this issue and know how to avoid it?
Thanks.
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Hi Michael
Even if you want to run in TAE buffer (because you need to recover the bands, etc), a test in TBE should help you clarify whether thermal diffusion (e.g., too high voltage) is a problem. Depending on the amount of available buffer in the tanks, most of the cations in the upper part of the gel can move to the lower side of the gel, thus making the intensity to grow up  (and the heat too, in order to maintain the voltage).
You could also re-circulate your buffer.