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Very introductory online source of information in evolutionary biology
We receive quite a lot of questions from layman in evolutionary biology in this site that are sometimes difficult to answer just because there is way too much to say. Why don't human keep evolving? is a perfect example. I often want to give some pieces of information and link to a good source of information for them to get introductory knowledge in evolutionary biology.
Obviously nobody that ask a question on a Stack website want to read an answer of the kind You should read that book!. It is often not really pleasant to read a book, it takes time and the OP that get this answer will likely just forget about their question.
What are the best online resources that offer a very introductory understanding in evolutionary biology?
Either a written course of videos such as Khan Academy videos would do (I haven't watched the evolutionary biology section of Khan Academy though and can't really judge of how good it is).

At least online, I think the single best introductory evolution resource is the Evolution 101 tutorial at UC Berkeley's Understanding Evolution project. The site has been designed by some of the top evolutionary biologists and evolution educators in the country, and does a very good job presenting a basic overview of how evolution works.

Thank you +1 I updated my answer there following your recommendations.

Perhaps an even bigger problem is that evolution questions provoke lots of answers from people who think they understand evolution but do not. Worse still, these answers sometimes get upvoted by people who share similar misperceptions.  In a perfect world we might be able to move toward the norm established on other SE sites: people don't simply make things up if they don't know enough to answer authoritatively. In the meantime, I'm not sure quite what to do beyond posting educational links in response to answers as well as to questions.

I like this resource very much. Frankly, I'm not knowledgeable about evolution (although it seems many questioners here have an even foggier image than I do) so I'm reading it for myself and finding it valuable. Now that the site has graduated, I think we should make it a community ad.

@rumtscho What a great idea! Do you want to start a meta thread about it?

@Corvus there is a thread for community ads, but I don't have the time right now to design one. Will see if I can do it when I come back from vacation.


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Why do adults lose hearing at high frequencies?
Children and teens can hear high pitched sounds that adults can't hear anymore. Why do adults lose high-frequency hearing?

Hearing declines with age and, typically, high frequencies are affected first. Age-related hearing loss (or presbyacusis (Kujawa & Liberman, 2006)) is progressive and starts at the highest frequencies and as a person ages the lower frequencies are affected. The reduction in hearing sensitivity is caused by the loss of hair cells. Hair cells are sensory cells in the inner ear that convert acoustic vibrations into electrical signals. These signals are transmitted to the brain via the auditory nerve.
Because of the tonotopic organization of the inner ear (the cochlea) one can conclude that presbyacusis develops from the basal to apical region. The following schematic of the cochlea from Encyclopædia Britannica, Inc. nicely shows the cochlear tonotopy (i.e., high frequencies in the base, low frequencies in the apex);

As to your question why the basal region is more sensitive to aging - that is a very good and interesting question that has not been answered, as far as I am aware. What is known, however, is that the basal regions of the cochlea are also more sensitive to the damaging effects of ototoxic antibiotics such as kanamycin (Stronks et al., 2011) and other aminoglycosides, and some chemotherapeutics such as cisplatin (Schacht et al., 2012). In low doses these compounds specifically target the basal regions over the apical ones (at high doses the whole cochlea is affected).
Perhaps the specific anatomy and circulation causes the basal region to be affected more by these chemicals. For example, blood flow may be enhanced basally whereby environmental and food-based chemicals may enter the basal region more readily (educated guesses, I can dig up references of authors posing these possibilities if necessary). This scenario may also contribute to presbyacusis, since it is quite imaginable that years of exposure to various substances may cause accumulation damage specifically to the basal parts of the cochlea.
In addition, since high frequencies are coded in the basal region, and given that sound enters the cochlea in the basal region (see picture below), all sounds, including damaging loud low-frequency stimuli first travel to the base on their way to their point of processing.

Because sound is analyzed as a travelling wave, sound travel stops at that point where the frequency matches the cochlear characteristic frequency. There the wave quickly dies out due to cochlear filtering (Mather, 2006); see the first picture above. In effect, more apical parts of the cochlea are exposed less frequently to (damaging) sound stimuli, while the base of the cochlea is exposed to all sounds entering the inner ear.
Hence, chemical exposure and acoustic wear and tear may be more prominent in the cochlear base, which can explain why presbyacusis starts at the very high frequencies and progresses to low frequencies (upward in the cochlea) from there.
References
- Encyclopedia Brittanica* The-analysis-of-sound-frequencies-by-the-basilar-membrane
- Kujawa & Liberman, J Neurosci 2006; 26:2115-23
- Mather, Foundations of Perception, 2009
- Schacht et al, Anat Rec Hoboken 2013; 295:1837-50
- Stronks et al, Hear Res 2011; 272:95-107

Well researched! Great answer!

@CRags - Thanks! The topic is closely related to my Phd :)

Love this answer, "whereby environmental and food-based chemicals may enter the basal region more readily."  It appears you are suggesting there is a relationship between premature loss of hearing and toxin intake.  I have long been an advocate of healthy, toxin-free living.  Thanks for sharing.

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how to find accurately the closest species to my plant species?
I'm working on some bioinformatic analysis of a non-model woody plant within the family with least information. So, I'm looking for a closest species to my plant. Any suggestion would be highly appreciated. Also, It will be great if I have know about the divergence time (? million year) between two species. Is there any documents or website to address this issue? Thanks

Please [edit] your question and explain what data you have. What species is this?  Do you have its entire genome? Specific genes? Are there any sequences for this species or no sequences at all?

If you know the species name, you can search TreeBase to find previously published phylogenetic trees which will point you towards close relatives to your species. Depending on how obscure your taxon is, though, you might not find much.

You can try http://timetree.org/ to get the divergence time if you already have the names of the taxa.

Maybe you could get some information from the Angiosperm Phylogeny Group:
http://www.mobot.org/MOBOT/research/APweb/
They even have a tree where model organisms are highlighted:
http://www.mobot.org/MOBOT/research/APweb/trees/modeltreemap.html

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How to get phased haplotype from vcf file?
Although I'm reading all days and nights, still confused in this area. I have hundreds of BAM files (from whole-genome sequencing of a given human population). I was wondering how I can obtain phased haplotype from raw vcf file? I read also about GATK, but didn't find a clear answer to my question. Please kindly tell me if this tool is suitable for this goal?
Would you please introduce me some practical guides and pipelines that explain such an analysis step-by-step?
Thank you for your help

What format are you hoping to end up with? Note that your question would be a better fit on bioinformatics.SE.

Thanks. vcf file format. Sorry, what's bioinformatics.SE, is the bioinformatic forum?

See bioinformatics.stackExchange for your future questions of the kind.

You should be able reformat BAM files to VCF files with PGDSpider.
I find using specific softwares for this type data formatting tends to be error prone and not practical when having a large number of files to reformat, so I personally like to simply read about the format standard and write the code to reformat myself but that's very much a matter of personal preference.

Thank you for your response and forum. However, variant calling from bam file is a long story, beyond just file conversion.

You can use either HapCut2, WhatsHap or my package.

There are multilple packages to achieve this. I have personally used bcftools (SAMtools). Here is a non exhaustive list:

SAMtools (bcftools);
VarScan2;
FreeBayes;
Platypus;
Deep variant

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How realistic is to use DNA for long term storage?
This is mainly a followup question to the recent paper Next-Generation Digital Information Storage in DNA.
Personally, while I agree about the data density of the format, I can't help point out the large issues with the inability to rewrite and recopy the data efficiently and potential issues regarding the "immutability" of the data. I'm curious about other takes on the publication and whether or not the stability of the DNA as well as the "writing" inefficiencies are better than I'm believing.

http://the-scientist.com/2012/08/16/dna-data-storage/, for the lazier.

My own view is that while reading DNA has advanced (and is still advancing) immensely, but writing is very much a work in process and still has a long way to go. As for stability/fidelity, I don't think there is a problem, considering that the Neanderthal genome was sequenced and it was stored in less-than-perfect conditions.

I can at least answer the question about the stability of the stored information. With my colleagues, I worked for several years to estimate the rate of DNA degradation at room temperatures. Our results showed that the information could  be retrieved after 100,000 years storage.

Did you publish the research?  Could you link to it?

Hi Jerome. Given the source of the studies, it's probably not published.


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Functions of tracrRNA and crRNA in the CRISPR/Cas9 system
I need some help about Crispr / Cas9.
The CRISPR/Cas9 technic consists (for bacterias) in "cut" bacteriophage's DNA, in order to make it unfunctional.
When the RNA is transcribed (RNA which is complementary to the bacteriophage's DNA), it will be associated with crRNA and tracrRNA to form the sgRNA (single guide RNA).
But I would like to know where crRNA and tracrRNA are from? and what are their function(s)/ goal(s).
Sorry If I made mistakes I'm French.

Note that the sgRNA is not present in the natural CIRSPR system and is only used in engineering as a replacement for both the crRNA and the tracrRNA.

I have also been wondering the same thing for a while now and I think that the best answer that I could find after reading several review papers on CRISPR/Cas9 and online information, points to tracrRNA as serving the role of multiplexing with pre-crRNA which helps pre-crRNA mature to crRNA. This process activates the crRNA. I'm not exactly sure how this multiplexing action "matures" the pre-crRNA or what exactly that even means, but that is as much insight as the literature provides (see e.g. this abstract to a conference talk).
There most likely are some more important functions that have not yet been discovered for tracrRNA yet.

Welcome to Biology.SE and thank you for your answer. Could you please also add the references to the review papers and online information you are mentioning? Thanks!


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Do bacteria with flagella self-rotate?
The bacterial flagellum uses rotation for generating forward motion, just like a helicopter main rotor does. Helicopters must utilize a secondary rotor to avoid self-rotation due to preservation of angular momentum. Alternatively they can have multiple rotors rotating at opposite directions. But all flagella rotate in the same direction.  Does this mean that the bacterial cell rotates in the opposite direction?
If so, what is the typical frequency of such rotation? Shouldn't the rotation disturb sensing and nevigation in the environment by the cell?

Welcome to Biology.SE! What research have you done before asking it here?

Google, and a bit of common sense. But no definitive conclusion...

My opinion as a physicist would be that yes, they do.
Assuming a sperm-like structure with a single flagellum at the centre of a long, thin cell of uniform density. They would rotate around the axis of travel.
However, for other cells, this could be mitigated in several ways:

If the cell was submersed in or floating on a liquid, it is likely that the effects of Archimede's Principal would be sufficient to keep it in a fixed orientation (this is how ships and submarines stay upright despite rotors that all go in the same direction - the least dense parts of a body always attempt to restore themselves to the highest position).
Fluid in the cytoplasm could act as ballast, providing a counterrotation force thanks to the gravitational effect on it.
If the cell happened to be particularily masseous in relation to the force the flaggela were providing, this would make any rotational force negligible.
If the flagella were located away from the axis of travel (through the centre of gravity - ie to the sides) the torque would have less of an effect.

As for the navigational aspect of your question, if these cells do auto-rotate, they will have done so throughout their evoloutionary histories and therefore in all likelihood will have evolved navigation methods capable of dealing with this.
I will however put a disclaimer on this, I am not a biologist and am weighing in from the point of view of a physicist.

1+2 are indeed nice ways to stabilize the cell. Has any such mechanism  been found?

3 sounds wrong. if the cell is too massive in relation to the force, then the flagella will not be useful for motility. I would say that if the flagella applies a forward force F on the cell, it must generate torque of approximately F*r, where r is the radius of the flagellum helix, probably ~1 micron. Thus, with similar approximations I would say that using flagellum to move in velocity of 100 body lengths per second must compel auto-rotation with frequency of about 100 Hz. This is reasonable as it is a typical rotation frequency of flagella.

@UriM, I agree, the flagella wouldn't be much use in that situation, but the cell would still be fairly stable

As for whether or not any such mechanism has been found, 1 + 2 must be true for any cell of non-uniform density with a non-rigidly located cytoplasm

point 4 applies to E. coli which has peritrichous flagella i.e. they arise at points over the entire cell surface and coalesce to form a flagellar bundle. There is a flexible component, the hook,  between the motor and the filament which enables this.

Yes, the bacterial cell body rotates, but more slowly than you might think.
Bacteria are examples of 'life at low Reynold's number'. Here are some extracts from  Howard C. Berg Random Walks in Biology (1993) Princeton University Press pp 75-80.

The Reynolds number is a dimensionless parameter in the equations of motion of a fluid that indicates the relative size of terms that describe intertial forces (forces required to accelerate masses) and viscous forces (forces due to viscous shear).

For a fish R is approximately 105; for a bacterium it is approximately 10-5. This difference has a profound effect on the physics of swimming.

The fish propels itself by accelerating water, the bacterium by using viscous shear.

To illustrate the lack of inertial effects at low Reynolds number, Berg estimates that if a bacterium stops swimming it comes to rest within about 1 µs and coasts for approximately 0.4 nm.

Flagellated bacteria swim by rotating one or more thin helical filaments... Torque generated by rotation of the filaments is balanced by viscous drag due to the counter-rotation of the cell and thrust generated by rotation of the filaments is balanced by viscous drag due to translation of the body of the cell.

In a recent paper from Berg's group direct measurements of the rotation of the flagellar filament and the cell body (for E. coli) are presented: the filament rotates at approximately 100 Hz; the cell body at 20 Hz.
The supplementary question is: Shouldn't the rotation disturb sensing and nevigation in the environment by the cell? Bacterial chemotaxis, at least in E. coli, relies upon time-averaged sensing of the concentrations of attractants in the surroundings via ligand-receptor interactions. Rotation of the cell body will not have any effect on these interactions.


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Can some colours be processed faster/more accurately than others?
Is the human visual / motor system able to track, and move in response to, objects of certain colours more quickly and reliably than for others?  By more reliably, I mean with greater accuracy in judging position and velocity, and with greater accuracy when moving in response.
Which colours are "most visible" in that sense, as a function of background and illumination?
This interests me specifically in the context of juggling: I like to juggle, and I wonder whether there's a difference in my response time and accuracy with different colours of balls/clubs.  Usually I'm indoors, and ceilings are usually white, sometimes black or brown, with strip lighting.  My only personal observations are rather obvious: dark colours can be hard to see against dark backgrounds, and white can be hard to see against white.


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Why is free ribose not reduced to deoxyribose rather than the reduction occuring on ribonucleotides
I cannot understand why deoxyribonucleotides are not synthesized directly from deoxyribose, but ribonucleotides have to be synthesized first, and only then can deoxyribonucleotides be synthesized.

Clarification
I assume that question is posed in the knowledge that ribose is an intermediate in the dark reaction of photosynthesis and can be synthesized in the hexose monophosphate shunt in animals. Hence I assume that the question is not why ribose precedes deoxyribose in the metabolism of mammals and other living things. Rather, the question would seem to be why ribose is not converted directly to dexoyribose, but only after ribonucleotides have been formed (in the ribonucleotide reductase reaction).
Possible Answer
Question as to why a metabolic pathway has one form rather than another can sometimes have a chemical explanation, but in this case I suggest answers can only be speculative.
Thus, one possible answer could be:

There is no need for free deoxyribose in the cell, and that if free
  deoxyribose were produced there would be potential competitive
  inhibition of enzymes that catalyse reactions involving ribose.

If an ‘RNA world’ preceded the modern ‘DNA world’, then one could argue that:

The simplest way for the shift from an RNA genome to a DNA genome would by the introduction of a
  specific reaction for the reduction of the ribose in nucleotides. The
  fact that the base that differentiates DNA from RNA — thymine —
  is also synthesized as a nucleotide (see my answer to this
  question)
  is consistent with (but does not, of course, prove) this idea.


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How can P450 distinguish between foreign and native compounds?
It is my understanding that P450 enzymes are capable of selectively degrading compounds that enter the cell from the outside (e.g. synthetic drugs) without damaging compounds that are metabolic intermediates normally found in cells. What is the mechanism that provides these enzymes with the ability to distinguish the foreign compounds that are substrates from the normal metabolic intermediates that are not substrates, if they are structually similiar?

False Premise
The false premise in this question lies in the end of the last sentence:

…the ability to distinguish the foreign compounds that are substrates
  from the normal metabolic intermediates that are not substrates,
  if they are structurally similar

The assumption here would seem to be that the similarity between normal metabolic intermediates and foreign compounds is too great for discrimination to occur. There is no reason to assume this.
Explanation

The substrate specificity of those P450 enzymes that metabolise
  xenobiotics is such that they will not metabolize normal cellular
  constituents.

There is a wealth of evidence that enzymes can have substrate specificity that is either extremely narrow or relatively wide. This obviously depends on the structure of the substrate binding site — its shape and size and the chemical nature of the amino acid residues there. Thinking in these terms one can envisage an enzyme that will discriminate against substrates with a single substituent at a particular position, but metabolize a variety of similar compounds that lack this. The evolution of P450 enzymes can be imagined as a process whereby mutations that allowed metabolism of toxic environmental compounds would convey an advantage and be selected for, but mutations that disrupted normal metabolism would be lethal (or disadvantageous) and be selected against.
The Broader context of P450 Enzymes
The question is posed in terms that might suggest that the only function of P450 enzymes is in removal of xenobiotics, and perhaps that they are only a feature higher organisms. In fact neither of these are true. Details of their occurrence in bacteria can be found in the Wikipedia article. However, a paper which I, as a non-expert, found interesting in this respect was published by Kawashima and Satta in PLOS ONE in 2014. This makes the point that there are two classes of P450 genes — those involved in normal cellular biosynthesis (e.g. steroids, cholesterols, vitamin D3 and bile acids), and those active against xenobiotics (e.g. plant alkaloids, aromatic compounds and fatty acids). Comparison of the genes for these in a variety of (mainly) vertebrates implies those involved in the metabolism of xenobiotics evolved from those involved in normal cellular metabolism, and such detoxification genes have arisen not just once, but on several occasions.


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What causes sodium channels to open?
What triggers the opening of sodium channels in a neuronal membrane? Is it acetylcholine that activates sodium channels in the postsynaptic membrane?
Are sodium channels like receptors that have to bind to something (like a protein or an ion) to open? If so, what is the substance that forces Na+ channels to open?

Sodium channels are primarily voltage-gated - these are the channels responsible for action potentials.
Many other receptors are ligand-gated, and these are typically the signal that causes the initial voltage change that opens the voltage-gated sodium channels; however, these channels are less selective cation channels and are permeable to ions like potassium as well as sodium. Still, their permeability to sodium is quite important and so they may also be thought of as sodium channels sometimes.
These include neurotransmitter gated channels like nicotinic acetylcholine receptors and AMPA (glutamate) receptors, and transient receptor potential channels like the TRPV1 receptor that is sensitive to painful heat and the chemical capsaicin which makes chili peppers "hot."

Thanks for the answer but I'm still confused. The Wikipedia link says that a change in the cell's membrane causes a change in the state of nearby Na+ channels and opens them. But I thought that a change in the membrane's electric potential (i.e. action potential?) is possible after the sodium channels have opened already and not before it. So, how can the cell's membrane potential change if no sodium ions have entered the cell yet? Do you see what I'm trying to say? I'm afraid that I'm not explaining myself well.

@stressedout Action potentials are driven by a positive feedback loop among voltage-gated sodium channels once a cell is depolarized enough. The initial depolarization is not an action potential and comes from someplace else, usually the ligand-gated channels I described. I recommend a basic textbook on neurobiology like Purves' "Neuroscience" or Kandel's "Principles of Neural Science" - both will explain this in detail.


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Separating DNA Fragments by Gel Electrophoresis. Are all the strands for one size the same?
My apologies if my question is too basic, and please point me to a more appropriate forum.  I am reading the textbook "Essential Cell Biology" by Alberts et al, and am consulting other sources as well.  I am reading Chapter 10 about DNA technology and have read how DNA is separated into strands of different lengths by restriction enzymes.  The different size strands can then by separated by Gel Electrophoresis.  I believe I understand these basics. My question is:
Do all strands of the same length have the same sequence of base pairs?  I wouldn't think      so, and this would be a problem in the next step, determining the sequence of one size fragment by  using special base pairs that are lacking a hydroxyl group. Isn't it possible I could have two strands that are 50 base pairs long (for example) that have the same beginning and same end and are thus cut by the restriction enzymes to a length of 50, but are most assuredly not the same within their sequence
The text says that once the different size strands are separated by electrophoresis, you can just cut out one of the bands and work with that.  That makes sense if you are guaranteed to only have 1 strand at that level of the gel agar or if the multiple strands are all the same exact sequence.
If you read this far, thank you.  I think more generally I need to know how many of these segments are produced when you use restriction enzymes and how long the average one is. And is there any guarantee for one chromosone (or all 46 chrmosones) having all the segments be unique sizes (I wouldn't think so)?
And even more generally :), I need additional sources that explain DNA sequencing in simple to understand terms.
Thanks,
Dave

You are correct: strands of the same length do not necessarily have the same sequence. The separation of DNA in gel electrophoresis is purely based on the hydrodynamic size.
The number of fragments produced by restriction enzyme digestion depends on the number of sites for that particular enzyme in your input DNA.  Restriction enzymes only cut at specific nucleotide sequences.  There is no "typical length" of fragment produced for this reason.
The idea of separating DNA on the basis of an RE digest is that a mutation that changes the sequence where the RE binds and cuts will reduce the number of fragments because the RE can no longer bind and cut at that site: 2 smaller fragments will now appear as one large fragment.  It will be visible on the gel because of the difference in number and size of fragments.
Typically, when running a gel, you are using DNA that has been selectively amplified to bias the readout to a particular (known) region of the chromosome. This electrophoretic approach works because you're not looking at all 46 chromosomes at once.
As far as sequencing is concerned, I'd recommend starting with Wikipedia.  Sequencing techniques are changing rather rapidly, so there's not one good source of which I am aware. To begin with, you can read about Sanger sequencing method. If you understand PCR then you won't find it hard to understand.

Thank you for your quick response.  I'm still not sure I have an answer to my question.  Perhaps the question is just too obvious and the answer is in front of my face.  If as you say, "strands of the same length do not necessarily have the same sequence" how does one separate them for subsequent analysis to specify that actual base pair pattern? If I understand the book correctly, strands will fall in the gel according to the number of base pairs they have.  Thus two completely different strands that are both 50 base pairs long will fall to the same height.

If I then try to figure out the sequence of this "strand", that is really two strands (or three, four, etc.) won't I just have a mess?

Yes, differentiating the strands is not possible on this basis: restriction enzymes can only be used to look at changes at a particular site, but it doesn't tell you what else is going on, or what the change was in the first place.  The sequencing method that you want to take a look at is Sanger sequencing...things have progressed since, but that is how you'd actually use electrophoresis to sequence.

All is dependent on how much you are sure about your experimental result. If you are very sure as in case of a plasmid digestion or a gene specific PCR then the band that you are getting will most likely be the strand of your interest.
However, when you are unsure about the experimental outcome then there is no better way to check what a band is composed of, other than sequencing. [in most next-gen sequencing experiments, the "libraries" are comprised of bands of sheared DNA migrating together, lets say at around 200bp. In this case the aim is to sequence the unknown]

The answer to the part of your question concerned with average lengths of restriction fragments is: if the DNA molecule being digested is of random sequence, and is 50% GC/ 50% AT, then the probability of finding any given short sequence at any position is 1/4N where N is the length of the short sequence. So, for example for a restriction enzyme with a 4-base recognition sequence such as AluI (AGCT) the probability is 1/256 and so the average length of an AluI fragment would be 256 bp. For an enzyme with a 6-base recognition sequence such as HindIII (AAGCTT) the corresponding number is 4096 bp. Please note that this is the average length, and is not an 'expected' length.
The answer to the part of your question about fragments of the same size co-migrating, how would you purify them on a gel for further analysis is trickier. I think the first thing to say is that you wouldn't do this for sequencing, and this is reflected in the various answers you have already received - sequencing no longer requires this sort of approach.
If you were forced into this position for some reason then I can think of two things that could help: the first option would be to run the analysis on an acrylamide gel when you can get much better resolution of different fragments (all 50 bp fragments are not the same molecular weight), but this is really only useful for small fragments (100 bp or less). However the most likely way out of the co-migration problem would be to purify the two comigrating fragments as a mixture and then ligate them into a vector whereupon each recombinant would carry one or other of the two fragments. This is really going back to the original reason why molecular cloning revolutionised things - it allows the 'biological' purification of DNA fragments followed by production of these fragments in large amounts.
You mention "special base pairs that are lacking a hydroxyl group" so clearly you are thinking in terms of Sanger (dideoxy-) sequencing, in which case you could clone directly into an M13 phage vector so that you could make ssDNA for direct sequencing. By sequencing a number of clones you would find two sequences, one for each fragment. You would still have to find a way to decide which one was the fragment that you were actually interested in, however.
Like I say, things just aren't done in this way anymore.

As I understand, you are confused how one would be sure that a band in a gel is the sequence one is looking for, seeing as sequences can be different and still have the same length (e.g. ACTTAT is the same length as GGCCGT but entirely different). So your actual question might rather be something like "How does one ascertain that a gel band contains only the sequence one is looking for?"
It is true that in practice, restricting a long sequence can lead to similarly long segments which are not the same sequence. It is indeed possible to cut out a small square from the gel which contains the band, and then dissolve the gel and extract the genetic material. Sequencing the result would give you one of two outcomes: 1) If the band contained only your desired sequence, your sequencing technique should render exactly that sequence as a result. 2) If the band contained more than one different sequences of the same length, your sequencing method will probably leave you with ambiguous results.
So sequencing would probably only confirm whether or not the band contained a single distinct sequence. It would not be useful as a method of separating the sequence you want for future experiments from any other sequence of the same length. One method to do this could be PCR (polymerase chain reaction), a procedure that produces vast numbers (billions) of copies of a specific target sequence. If a PCR was carried out to amplify the target sequence contained in the gel band, any other sequences of the same length would not have any copies produced. They would still be present, but being vastly outnumbered by the sequence relevant for your experiments should prevent the other sequences from having any significant effect.
However, as dd3 noted, you would not normally run a the result of a restriction on a gel and then try to separate your target sequence from the rest. Rather, you would simply use the PCR method described above directly after using the restriction enzymes, and then run the result on a gel. You would then get a very strong band for the sequence you were looking for and much weaker bands for everything else.


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what the best E-value cutoff in the miRNA homology search
I'm trying to use miRBase database to annotate some microRNA sequences in length 22-24bp. I highly appreciate if someone let me know whether I should do this BLAST against mature miRNA or stem-loop sequence database? and the other main question is that which number of E-value is reasonably accepted as a lowest E-Value in the miRNA BLAST results? Also, what tool you suggest to perform miRNA enrichment analysis? for example we can use DAVID for gene (coding sequence) enrichment analysis. Using such analysis we are able to assign probable functions and biological roles to a group of miRNA (not only one miRNA). Any comments and idea is welcomed.

If you are using BLAST for short nucleotide searches then you should reduce your word size to <=7. It is better to align with mature sequences.
E-value depends on the database size. If your target database is huge then you should increase the e-value cutoff. If miRBase (for a few organisms) is your database then there is no need to run BLAST, also. You can simply run SSEARCH (Smith-Waterman Local alignment algorithm).
If you are running BLAST against miRbase you can set the e-value cutoff at 10. This is fine because there are many similar looking miRNAs with different function. If you reduce the e-value cutoff then you may lose these.
There are some other criteria for miRNA conservation- e.g there should be very high (almost exact) similarity at the seed region. After the BLAST, make sure that seed region is conserved.

Can we can use DAVID for miRNA functional analysis.

I don't think DAVID can be directly used (miRNAs have to be annotated in their database). What you can do is that you can predict the targets of your miRNA and then pass them to DAVID. DAVID won't do the predictions but there are other programs (see this post). You can also try BioMart (provided by ENSEMBL) for getting GO details. Even BioMart does not provide functional annotation for miRNAs (I just tried, to verify). So best thing is to check for the targets.

Thanks for your feedback. I want to use miRBase to do this blast that its default word size is 4, is nice for short sequence homology search, but the E-value cutoff is 10 by default that seems a bit high for miRNA (short sequence) in my opinion since the probability of having good alinment by chance is increased as the sequence length is decreased, is it right or no?

Yes that's true. Also that probability increases when the database size increases. If you find the answer suitable then you can accept it (A tick mark on the left).

Ok. Regarding what we discuses, could you please suggest me what is the best E-value cutoff in the case of miRNA homology search? I'm a bit confused about the selection this parameter.

Dear friend, as you said, DAVID can not be directly used for miRNA analysis. Also, miRNA target analysis using this tool is a different story. at the moment My focus is on miRNA not their targets.

@Mary- edited the answer. No you cannot directly annotate miRNA. You need to find its targets first. E-value of 10 is fine (don't go below this).

Thanks a lot for being with me! what about the mature miRNA and stem-loop sequence database, both available in miRBase! which one should I choose as a database? my concern about this topic is that I should confirmed the best hit of the blast with what I looking for it in the laboratory.

I mentioned in the answer- align against mature. Other regions are not that well conserved.

10? Really? I haven't worked with miRs but 10 seems extremely high. In fact, working on short nucleotide sequences, I wouldn't recommend using the evalue at all. I'd go for plain %id and % coverage instead.

@terdon 10 is safe. I think the online BLAST asks for evalue cutoff. It assigns 10 by default for short nucleotide searches. But for miRNA, I think more parameters should be defined. I think gap start penalty should be high (5` region should match perfectly). Using a short read aligner like bowtie is also not a bad idea. It allows you to set a seed sequence and lets you define allowed mismatches in seed and non seed regions separately.

Absolutely. Your parameters sound fine. I just wouldn't use an evalue of 10 as a way of determining whether a hit was interesting. I'd leave it at 10 or higher to be sure to get everything and then ignore it and filter on other things as you suggest.


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Why does the ecosystem of my house seem inverted?
My understanding of a typical ecosystem is that it is like a pyramid, with the mass of prey at the base and the apex predators at the peak.
However, the arthropod ecosystem in my house seems to be the inversion of this.  When I see critters, I mostly see spiders and centipedes, the apex predators.  Rarely do I see anything else.  This seems upside-down.  Shouldn't the apex predators of an ecosystem be relatively rare?

I’m curious why you think the apex predators should lay low. They’re apex predators. If anything should lay low, shouldn’t it be the prey? Also, there are different ways to remain undetected.

@anongoodnurse – I guess I'm thinking of something like a jaguar or panther which is very elusive.

Spiders that use webs are quite easy to spot for humans because they are stationary and they tend to be in the same places. The same isn't true for their preys which, for the record, need not be part of the ecosystem of your house: they can be some animals that are passing by.


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How do fatty acid and polyketide synthases handle the varying size of their substrates?
Polyketide synthesis involves six modules, each containing certain common constituent enzyme activities (KS, AT, KR) that are involved in extending the chain (see e.g. Dutta et al.(2014) ). The growing polyketide, attached to acyl carrier protein, is passed on from one module to the next.
An analagous situation is seen in the single Fatty Acid Synthase complex. How can the same enzymes in later modules recognize what is a structurally different intermediate? Surely this contradicts the idea of substrate specificity of enzymes.

I've looked at the paper very quickly (I've other things to do this evening.). Am I correct in thinking that your question is as follows — restricting it, for the moment, to the polyketide synthase in the paper? "There are six modules in PK synthesis, each containing some common enzyme constituents (KS, AT, KR) that are involved in extending the chain. The growing PK, attached to acyl carrier protein, is passed on from one module to the next. How can the same enzyme in later modules recognize a different intermediate?" If that is correct I will rewrite the question for you (and answer it).

I had time to answer what I think is your question, so went ahead and modified it and answered it. Let me know if I have misinterpreted anything.

@David Yeah, this is the most correct interpretation of what I mean. Simply to say, I was asking how enzymes on later modules bind to large polyketides even the enzymes are as same as the previous ones and the enzymes on later modules should violate substrate specifity of enzymes.

Although the substrate is different at each stage of chain extension,
  the part involved in the reactions — and recognized by the enzyme — is
  the same. This is a common feature of polymerization reactions,
  whether the catalysis is by a single module (ribosomes, DNA
  polymerase) or multiple modules (polyketide and fatty acid synthesis).

This is illustrated by the modification made below to a part of Figure 1 of the paper cited in the question. An overlay in yellow has been made to indicate the common structural features of the polyketides of different length.

Of course, the rest of the chain may well interact with the enzymes, but as this is not at their active sites it can be in a less specific manner, perhaps in a ‘groove’ containing appropriate interating chemical groups.
The question of substrate specificity is not the point of interest in the paper on polyketide synthase where the focus is on the way the elongating substrate is passed from enzyme to enzyme and module to module (the resolution is also too low). It has, in any case, been addressed in previous work on mammalian fatty acyid synthase, where exactly the same single (dimeric) multidomain complex acts on each intermediate until the chain length is 16-18 C atoms. However the question of chain length is taken for granted and the discussion is about discrimination between chemically different intermediates. Thus in a 2008 paper by Maier et al. a substrate tunnel in the ketoacyl synthase (KS) module of the fatty acid synthase (mFAS) is compared with that in polyketide synthase (PKS):

“Despite the pronounced structural similarity between the KS domains of mFAS and PKS, the selectivity of FAS for saturated acyl chains can be explained by a considerable constriction at the base of the active site phosphopantetheine binding pocket leading into the large acyl chain substrate binding tunnel, which connects both active sites of the KS dimer (Fig. 4, A and B). A number of residues lining this narrow tunnel are highly conserved in mFAS but are substituted with smaller residues in all DEBS (6-deoxyerythronolide B synthase) PKS modules (Fig. 4B), resulting in a wider, more permissive tunnel in the KS of DEBS.”

It is important to think of substrate specificity in molecular terms — e.g. what part of the substrate binds to the enzyme and how specific has the binding to be. In this case the substrate is attached to the long phosphopantetheine chain of acyl carrier protein, so that recognition of that and the part of the fatty acid or polyketide attached to the sulphur may be enough to discriminate against ‘free’ extraneous molecules. And a tunnel is permissive as a binding site: if the chemistry and radius are right, there is no need for the whole of the tunnel to be occupied.
Tunnels of various types are found in other enzymes or organelles involved in elongating molecular ‘chains’, e.g. polypeptides and nucleic acids. The tunnel on the ribosome and that through which RNA or DNA passes has to accomodate chemical disparate molecules (four different bases, 20 different amino acids).

I get it. So the rest of the chain is made to be not on their active sites by its enzyme mechanism and the enzymes recognize the yellow parts, thus the enzymes can do their reactions regioselectively. Perhaps, can you locate me which paragraph on the reference that tells about these active sites?

I have modified my answer to deal with the point you raise, as I think that more generally useful than a conversation in the comments. I'm not sure what level you are at, but if you can get hold of the Science paper I reference and have the time and background to work through it, it will give you a better idea of active sites and substrate binding sites.


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Are mitochondrial genes decoded in the same way as nuclear genes?
Mammalian mitochondrial genomes contain only 22 tRNA-coding genes, which is an insufficient number to decode mRNAs under the standard wobble rules.
How is translation of mitochondrial mRNAs achieved with this number of tRNAs?
The possibilities that occur to me are that certain genes encode multiple tRNAs (but this appears unlikely as each gene has its own promoter) or that mitochondrial tRNAs can adopt alternative three-dimensional structures, each with a different anticodon loop and hence a different anticodon.

Answer Summary

Translation in the mitochondria of mammals differs markedly from that
  in the cytoplasm. Although it has more similarity to translation in
  prokaryotes, it also shows significant changes from the latter. A smaller
  repertoire of tRNAs is able to translate a ‘simplified’ genetic code
  through structural alterations and base modifications that allow their anticodons
  to decode groups of synonymous codons, not possible according to the standard wobble ‘rules’.

Relationship to the eubacterial translation system
One of the pieces of evidence for the endosymbiotic theory of the origin of mitochondria was the fact that their translation machinery was more similar to that of prokaryotes than to that in the eukaryotic cytoplasm. The similarities to prokaryotes included the (smaller) size of their ribosomes, the susceptibility of their ribosomes to antibiotics specific for bacterial ribosomes, and the formylation of their initiator tRNA — fmet-tRNA.
Although in no way rebutting the endosymbiotic theory (which is widely accepted), mitochondrial translation shows several differences from that in prokaryotes. These include there being a single tRNA to decode methionine in both initiation and elongation (discussed below), the lack of leader sequences preceding the initiation codon, hence precluding a Shine & Dalgarno type of recognition by the ribosome, and additional ribosomal proteins which are thought to “provide a specialized platform for the synthesis and membrane insertion of the highly hydrophobic protein components of the respiratory chain”.
Decoding with 22 mitochondrial tRNAs
General Review: Suzuki et al. (2011) Annual Review of Genetics 45, 299–329
As the question states, mammalian mitochondria encode 22 mitochondrial tRNAs, which would be insufficient to decode mRNAs using the standard genetic code and the wobble ‘rules’ that apply to translation in the eukaryotic cytoplasm and in prokaryotes.
One way out of the dilemma would be if some of the tRNAs were encoded in the nucleus and imported into the mitochondrion in a similar manner to that of nuclear-encoded mitochondrial proteins. Although this turns out to be true for certain organisms (although the import system is different from that for proteins), and important in some cases, this is not the answer for mammalian mitochondria. Thus, as reviewed in Current Genetics (2009), the mitochondrial tRNAs encoded in the nuclear genome of H.sapiens only duplicate the 22 mitochondrial tRNAs and their anticodons. (The one exception is an extra Gln(anticodon CUG) in addition to the Gln(anticodon UUG).)
Let us look more carefully at the problem. Although there are 61 codons specifying amino acids, because of wobble, fewer than this number of tRNAs are required to decode them. The left-hand frame of the figure below shows the standard genetic code, and the minimum number of tRNAs needed to decode it using the wobble ‘rules’ shown beneath, indicated by the colour-coded rectangles.
For those not familiar with the actual pattern of wobble (summarized in my answer to another question), in elongator tRNAs the 5′-position of the anticodon has some flexibility such that a G in that position can form base pairs with U or C in the 3′-position of the codon, an inosine (I) can pair with U,C or A, but an anticodon C can only pair with a codon G. The unmodified bases A and U are rarely found in anticodons, but a variety of different chemically modified Us are found (U* in the figure) which only pair with A. Thus, the minimum number of elongator tRNAs needed would be the number of coloured rectangles — 36 — and, as there are separate species of tRNAmet for initiation and elongation, the minimum number of tRNA species is 37.

When the human mitochondrial genome was sequenced, two anomalies stood together: the limited number of tRNA genes already mentioned and (more sensationally) changes from the standard genetic code (indicated in red on the right-hand side of the figure). It was immediately realized that these changes served to rationalize the code so that all synonymous codons were in groups of four or two. If it were assumed that the decoding specificity or wobble of mitochondrial tRNAs were different, then 22 tRNAs would be sufficient using the ‘wobble’ shown beneath the right-hand frame of the figure for groups of four amino acids (dark blue) and groups of two amino acids with codons ending in A or G (red).
One further requirement was that a single tRNAmet be involved in both elongation and initiation, a requirement the fulfilment of which has already been mentioned.
What features of the tRNAs could change the wobble ‘rules’ that have been conserved between prokaryotes and eukaryotic cytoplasm? There turn out to be two types of changes. The first, and most obvious, is the fact that certain (but not all) mitochondrial tRNAs have non-canonical cloverleaf structures, the most extreme being tRNAser in which the whole D-loop is missing. The second is the post-transcriptional modification of bases in the anticodon — just as is the case for tRNAs in the cytoplasm, this can markedly affect decoding specificity.
It is not yet possible to describe the codon–anticodon interaction in mitochondrial translation in the way that this has been done in for the cytoplasm or for prokaryotes. The problem would appear to be the difficulty of obtaining sufficient pure tRNAs. Whereas it is easy enough to clone and transcribe the tRNA genes, this is insufficient without the secondary chemical modifications of the bases which is key to the structure and function of the tRNAs.

Wow. Thanks for taking on the effort to write this! It’s been really helpful. May I ask a very dumb question? What do you mean by “the anticodon decoding both 3’A and G, or 3’ A,U,G and C.”?

@YamanKJ — I have revised my answer to explain the tRNA decoding better using a diagram of my own. I hope it's clear (and that there aren't any mistakes in the diagram.)

At first I thought that the "wobble pattern" was another name for the "Watson and Crick pattern" as I'm too naive with genetics, obviously. Your answer is indeed very informative, and you may have just made a frame-shift change (to use genetics' terms) to my level of comprehension of this topic. Every time I read your answer I understand one more detail. I think you need to edit "modified Us only pair with G (here I think it should be A instead of G?)."  and flip the colours here "groups of four amino acids (red) and groups of two amino acids with codons ending in A or G (dark blue)."

Forever grateful for this detailed explanation!

@YamanKJ — Thanks for pointing out the typos — one can go blind writing these things, especially when the output is out of view. Happy to help. I used to be more involved in protein synthesis, so I welcome the opportunity to check out how things have progressed in a particular sub-area. Of course, the danger is always that one gets an incomplete or distorted picture, but then one hopes someone will come along and point it out.

It turns out (and I’m not a 100% sure of this) that every tRNA-coding gene could give multiple variants of anticodons by different folding ways, so that you’d have enough anticodons in the end.

No, that's incorrect. tRNAs do not get their anticodon identity via folding. The anticodon is defined by the gene sequence.

@KonradRudolph That’s what I’ve been told when I asked my tutor. Do you have any idea about the answer to the aforementioned question?

Your tutor is definitely wrong (I should know: I literally wrote my PhD thesis on the topic). Unfortunately I don’t know the answer for mtDNA either but you forgot at least two possible explanations in your question’s list: (1) mtDNA only uses 22 different codons, (2) mtDNA co-opts nuclear tRNA for its translation. I suspect that the answer is a mix of these two explanations (i.e. mtDNA mostly uses a reduced pool of codons, and almost certainly uses nuclear tRNAs because these are abundant and the genetic code is almost the same as the nuclear one anyway, and translation happens in cytoplasm).

@KonradRudolph Well, I definitely trust Cambridge students more than I trust our university’s (he’s a great, great tutor nevertheless). I’d thought about what you’ve suggested beforehand but(1) is wrong because I’ve seen a table that contains every codon and its corresponding amino acid, they surely need more than 22 codons. (2) I thought it was not likely given the fact that mtDNA has 4 stop codons(only 2 of which are shared with nuclear DNA)so if the tRNA of nuclear DNA fits, the translation would not stop there. Could be that mtDNA shares SOME of the tRNA w the nuclear DNA. (2) makes sense!


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Can cellar spiders survive long immersion in water?
One morning i got into my tub to take a shower, i didnt check it this time and didnt notice a cellar spider that was in it (its pretty common for them to be in there i take them out all the time) and started showering. After a few seconds i noticed it floating in the water and hurried to get it out.
I got it out and at first it seemed like it was alive, its legs were holding onto my finger but it wasnt moving at all. When i managed to get it onto a dry surface it just curled up and it looked like it died. I left it on the washing machine, saddened by the fact i just killed it, and finished up my shower.
I forgot to take it away and left it on the washing machine. I remembered after a few hours that i left it there and when i went to check on it, it was gone. Nobody was in the bathroom except me during this time so i think it literally got up and left. Or some other bug carried it away but i highly doubt it.
So my question is did the spider actually survive? And is that even possible? It was floating in running water.

"...did the spider actually survive? And is that even possible?" Absolutely (unless someone else removed it, it's the only explanation possible.) Drowning a spider takes more than a few seconds, and it probably looked dead because it was weak/exhausted. It's nice that your response was to save it; it worked. :)

Yeah thank God I managed to do it hehe.

Who would have thought this question would get such a stellar answer? Thanks for asking it. My close vote has been retracted, and TIL.

The chitin exoskeleton of spiders is hydrophobic. That, coupled with their fine hairs, means water can´t usually get to their lungs/respiratory apparatus, as a layer of air will form around their whole body due to the surface tension of water. If however there are some surfactants breaking the surface tension, like soap or detergent, they can easily drown.
The spider however most likely played dead and just wandered away once she felt safe, or she really was suffering from oxygen deprivation. If so, it takes a while until their system starts functioning again.
Drowning would take couple of hours to days.
What is the Best Way to Kill a Spider?
Can Spiders Drown?
Submersion tolerance in a lakeshore population of Pardosa lapidicina (Araneae: Lycosidae)

Thanks for the information! The tub was clean and I didn't use soap yet so it probably did survive.

Hi and Welcome! This answer makes me very happy this site exists. Thank you so very much! (Also, this site requires sources which support the answer. I added those so as to meet that requirements.)

Nitpick: AFAIK spiders don't have lungs, they have spiracles or stigma connected to breathing tubes.

They have structures analogous to lungs, called book lungs, as they look like stacked pages of a book, where hemolymph runs through which increases surface area to blood ratio.


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How do temperature sensory neurons differentiate different temperatures
There are both cold and hot receptors of temperature which use TRP ion channels. When a certain temperature is achieved one of the TRP channel is opened and Ca2+ ions enter the first order neuron,
My question is that if several types of TRP ion channel are present on this neuron, how does the neuron differentiate mildly hot temperature from very hot temperature?

Welcome. Can you add the source of the figure you included? Do you have a clue what the answer might be to this question? I mean, there are differences between the receptor activations, so what do you think might be going on here? Please edit the question by including the source of the image and an attempt to your answer

I´d say that this is just for presentive purposes.
Heat activated neurons will mainly express one TRP-receptor type and show activity at this specific temperature range.
I did not find publications that show coexpression of all of those receptors except for TRPM8, TRPA1, TRPV 2 with TRPV1 ("TRPV1 was expressed in 29% of TRPM8-positive cells in dorsal root ganglion sections"-https://pubmed.ncbi.nlm.nih.gov/15050705/ , "TRPV1 is coexpressed on the vast majority of TRPA1-expressing sensory nerves "-https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3417484/ , "significant co-expression was seen in 20.7% of TRPV2-positive neurons"-https://journals.lww.com/neuroreport/abstract/2003/12020/co_expression_of_heat_sensitive_vanilloid_receptor.23.aspx  )
Here some quotes from another paper that would corroborate that:
"Some fibres respond to a moderate threshold (∼43°C), whereas a smaller percentage respond to high-threshold (∼52°C)"
"observed that cutaneous warm fibres of the hairy skin of primates could be divided into two groups"..."However, one group of neurons had an average maximum response at 41°C, whereas the second group continued to increase its firing rate above this temperature."
https://www.nature.com/articles/nrn1141
(But if it were the case that there were multiple receptors one hypothetical way of discriminating between different receptors would be the intensity of activation which can change the frequency of action potentials,that might be higher for one receptortype due to higher ion conductance,or all receptors together might be active at once at certain temperature)

I think this answer overemphasizes firing at a specific temperature, which isn't really how these neurons code. Your uncited parenthetical paragraph at the end is more representative.

This sentence is a contradiction: "I did not find publications that show coexpression of multiple of those receptors, though there are some that can occur together on the same neuron"


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What percentage of a Neanderthal's DNA could be in a Denisovan?
My questions arose from an online course video at 3:11-19
The professor mentioned that the Neanderthals were inbred with their cousins breeding together but the Denisovans were very diverse , with 70% Neanderthal's DNA, there was found mixing between Denisovans and an even earlier mysterious species.
The followings are my questions:

Did he mean that most of Neanderthals were prone to inbreeding? Why?
I'm not sure if that was exectly what he meant by 70% Neanderthal's DNA could exist in a Denisovan. If that were true, why not just called it Neanderthal if only 30% is not Neanderthal?

You should be very careful trying to conclude something from what is known about Neandertal genetics.
To answer question 1: NO! This is probably not what he meant. To date, we have only two high-quality full genomes of archaich humans, one being the (Neandertal from Denisova cave, called Altai Neandertal (Prüfer et al., 2014)) and one being the (Denisovan from Denisova cave (Meyer et. al., 2012)). From this sample (which is exactly one Neandertal) you cannot draw conclusion for the whole population. It seems, however, that this Neandertal individual from the Altai Mountains was indeed inbreed (high runs of homozygosity in the genome) - it does, however, not even tell us in what way her parents were related. Prüfer et al. (2014) write

We conclude that the parents of this Neandertal individual were either half-siblings who had a mother in common, double first cousins, an uncle and a niece, an aunt and a nephew, a grandfather and a granddaughter, or a grandmother and a grandson.

You cannot say more. You also cannot say if this was due to preference in mate choice, or caused by the fact that this group was very isolated, or  that this population had an extreme small effective population size (note that it is debated whether the last two effects are actually equivalent).
To make a long story short: based on the available data it is not wise to draw general conlcusions about Neandertal mating behaviour. Altai is the only example where direct inbreeding could be shown, all other genetic data from Neandertals suggest that genetic diversity was just low.
To answer question 2: I do not really know what this means, 70% Neandertal DNA in Densiovans. I can just generally tell you something about Densiovan-Neandertal differences. The Densiovan might be the only (at least mammal) species that was defined based on genetic information alone. Alle that was found were a few teeth and a pinky bone. People thought this was another Neandertal. After sequencing the DNA from that bones to high coverage (vaguely, a measure of qualtiy), one found that the DNA falls both outside the variation of modern humans - showing that this was not a Homo sapiens, but interestingly it also clearly falls outside the variation of known Neandertal sequences - showing that this was not a Neandertal. It also became clear that Densiovans are more closely related to Neandertals than to modern humans (you might now ask yourself how this is possible with just one genome. In fact, there are more genetic data availabe: low coverage genomes, exome sequences, mtDNA and all of them show the same thing: Neandertals and Densiovans are different but more closely related to each other than to modern humans). Overall, Denisovans also seem to have had rather low genetic diversity (which by the way is also a characteristic of modern humans).
Long story short: There seem to have been admixture events between Denisovans and Neandertals but I do not know how to relate this to the 70%. Prüfer et al. (2014) detected about 0.5% contribution from Neandertals in Denisovans.
One last comment about this mysterious species (nobody says it but most people think of Homo erectus): again, Prüfer et al. (2014) found some interesting patterns of derived allele sharing between present-day Africans, Denisovans and Neandertals (this should be equal for African-Denisovan and African-Neandertal comparisons, but present-day Africans share about 7% more derived alleles with Neandertals than with Densiovans - even 13-16% in alleles fixed in present-day Africans). This is either ancient population structure (however, with very deep lineages) or, as suggested in the paper, due to gene flow from a (genetically) unknown archaic hominin into Denisovans.
I hope this clears things up a bit. Feel free to ask in the comment and I will edit the post as it might be a bit unstructured.

Can you explain about what you mentioned --"Overall, Denisovans also seem to have had rather low genetic diversity (which by the way is also a characteristic of modern humans)"--     As what you said ,It seems both species have rather low genetic diversity, but with whom we are comparing ?  Does that mean there is a mistake in the video ,which mentioned -- the Denisovans were very diverse--?And also being against the quote from the link you give --several gene flow events occurred among Neanderthals, Denisovans and early modern humans--?

I compare genetic diversity between 'relevant' species that have a comparable evolutionary history and are closely related, i.e great apes: chimpanzees have about 3-4 times more genetic diversity than present-day modern humans (and other apes even more). I have not seen the whole video, so I am careful calling it a mistake. In the Meyer (2012) paper cited above you will find evidence that the genetic diversity in Denisovans was by far lower than that in present-day modern humans. And yes, there seem to have been multiple gene flow events.

Thanks for your reply ,but I still not understand :Why couldn't  "multiple gene flow events" relate to "high genetic  diversity'' ? Aren't they equivalent ?  To quote-- present-day Africans share about 7% more derived alleles with Neanderthals than with Densiovans--(I conclusively say that it means we are more relative to Neanderthals than Denisovans ) and-- It also became clear that Densiovans are more closely related to Neanderthals than to modern humans --Can I say that Neanderthals were tend to interbreed so their genetic diversity were higher than both modern humans and Denisovans?

You are mixing up some concepts there. 1. Gene flow does not automatically cause genetic diversity. If you have gene flow between hypothetical identical populations, the diversity stays constant. Neandertals and Denisovans were closely related - gene flow obviously did not lead to excess diversity in any of them as both populations lacked high levels of diversity. 2. The 7% shared derived alleles do not mean that Neandertals and humans are more closely related than humans and Denisovans - they just share some haplotypes. In fact, humans are equally related to Neandertals and Denisovans.

(continue) 3. You cannot say that Neandertals tend to interbreed (see my answer. The high quality genome available just happens to come from an inbreed individual. You will find inbred individuals in almost any population, also in present-day modern humans. You must not generalise this to the whole population.) 4. Inbreeding does not cause  genetic diversity. In contrast, inbreeding decreases genetic diversity as closely related individuals share alleles by descent so you will have a lot of homozygous sites in inbreed individuals (homzygosity is proxy for genetic diversity).


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Assays for protein or lipid content of insect pupae
Does anyone have any recommendations (based on personal experience) for rapid, cheap, accurate assays for estimating protein & lipid content of individual insect (mosquito) pupae?
So far I've come across chloroform/methanol extraction for lipids, and Kjedahl procedure + Nesslerization for protein (link).
Mosquito pupae are not particularly tough as insect pupae go, but ideally I'm looking for personal experience that the method can cope with the sorts of sample I need to use it for.
(I strongly suspect similar assays have been developed for Drosophila so have added that as a tag.)


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Antigen molecular mimicry
Let us consider a situation in which the body is attacked by a microbe, and  the microbe is captured by the immune system for recognition of surface antigens.
The surface antigen recognized mimics one of our self antigens.
It is stated in textbooks that this situation results in the immune system attacking  the microbe along with the tissue containing the self antigen(auto-immunity).
(Reference: Robbins and Cotran pathologic basis of disease)
If a microbial antigen mimicks self antigen, why is the microbial antigen attacked instead of being ignored in the way that all our self antigens are ignored by our immune system?

There are different type of immune cells which falls under either innate or adaptive system.
The cells under the innate are first to respond microbial attack and are able to detect them in extracellular environment by recognizing proteins/peptides called Pathogen-associated Molecular Patterns (PAMPs) on their surfaces; these are only found on microbes.
But when the innate system fails to clear the microbes in time, the adaptive  system is kicked-in. This involves T-lymphocytes and B-lymphocytes. T-lymphocytes are trained to ignore cells that are able demonstrated that they self but B-lymphocytes which produces antibodies/immunoglobulins to the detected antigens do not have a mechanism of ignoring self cells.
So the problem lies with when the immune response gets to the point where antibodies are produced, at that point produced antibodies simply attack without discrimination.
I hope this simplified version would help you.


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Why did a resazurin broth bottle turn pink immediately after being inoculated?
I have lawn cultures of aerobic bacteria kept at (2-8 °C), which I am trying to revive since they were kept longer than recommended. First, I tried to incubate a broth culture inoculated with a loopful from such cultures and then subculture the next day onto an agar medium. However, they showed no growth on the solid medium.
I resorted to another approach: I immersed a very heavy growth in a few mls of broth which was inoculated into manual blood culture bottles (rubber capped vials), as those:

Some of the bottles turned pink immediately after inoculation. Subsequently, I stained the immersion of these with simple stain (Safrinin), and it revealed the presence of the targeted organisms.
Is there any explanation for this other than that there were viable cells in the inoculum added to the blood culture bottles?
If there were viable cells, how come the color appeared too soon? Large number of viable cells maybe?
Did I push oxygen in with the inoculum through the syringe, so that the indicator got oxidized?

Some other manufacturers say this for the quality control part "color: Light straw coloured solution with upper 10% or less medium pink on standing", i saw some of the unused bottles have a slight pink color and I thought they were ill-manufactured. Maybe i should do a positive control and a real sterile control (inject sterile broth into a bottle, not just use a non used bottle), sorry I'm a dumb brick.

Definitely follow up with those controls, but very likely introduced oxygen when inoculating the tubes. Presumably there's oxygen in the broth and the syringe. I work with a lot of strict anaerobes, and I'm amazed at how sensitive resazurin indicator is. I can be pretty confident that my culture tubes were left open when cycling them into the anaerobic chamber, but sure enough there's always one or two where the anaerobe media goes pink as soon as I dispense it, just from that tiny bit of oxygenated atmosphere left in the tube.

@MikeyC they deleted my reply for unknown reasons. I wanted to say thank you and if this is not good to fit as an answer then what is!

Humble Mikey, i noticed that the "pink" i was refering to is the same color that appears when i injected mere air into a bottle, although i wasn't using aseptic manners, but if the indicator is too sensitive even to few oxygen, the few cells of air contaminants i might have introduced won't produce that few oxygen, right? Thanks.

i meant to say "..... when i injected intentionally mere air....."


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Wash adsorbed protein without destroying Biotin-Avidin-Mechanism
I am looking for reliable protocols to wash protein (e.g. Fibronectin) adsorbed onto glass surfaces. According to Protein immobilization literature, people usually use 1% SDS and some incubation time. For me this did not work.
Furthermore, the removal step should not destroy an existing avidin-biotin-mechanism.
Does anyone have experience with that? Or is there some kind of standard protocol, which I am missing?

More detailed process:
I have a quartz coverslip with covalently bound Biotin on some surface area. First I add Avidin (Sigma, 10µg/ml), then biotinylated human plasma Fibronectin (Sigma, 10µg/ml). Now I want to wash adsorbed Avidin/Fibronectin, so I have free "glass" area. This is important for me.
For some reason changing the system is not an option for me.

SDS shouldn't appreciably disrupt avidin-biotin interactions. The complex is extremely stable.

@canadianer SDS will disrupt the avidin-biotin bond at high enough concentrations. 1% is rather high (similar to Laemmli sample buffer), so that may be why the OP's experiment didn't work.

Can you explain what this did not work means in greater detail?

@MattDMo I have put streptavidin in SDS-PAGE loading buffer (2% SDS) and run it in an SDS-PAGE gel with only negligible denaturation. This paper has similar, and more formal, findings. I've also washed streptavidin conjugated chromatography columns with SDS without any leakage of the target protein. That said, it's hard to say if the SDS is or is not affecting the experiment. It would help if the OP gave the protocol and explained why it didn't work.

@canadianer that's really interesting. Did you add a reducing agent (DTT, BME) and/or boil the sample?

@MattDMo I don't remember if I had a reducing agent or not. No boiling though. Boiling + SDS = unfolding

@canadianer Let's continue this in the Biology chat room - it's kind of off-topic here.

@MattDMo Sure..

Sorry guys, time shift. I'll explain in more detail as soon as I'm back on the PC. Avidin biotin was totally stable with sds 1% (actually I used laemmli, 2hrs incubation at 4°C). But none of the adsorbed protein was washed away.

Do you want to wash off the fibronectin?

yes, except the avidin-linked fibronectins.


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Do snakes get dizzy?
I've seen snakes get captured in sacks and I've wondered if the snake would get dizzy if the sack were twirled around.  I have not done any tests (and don't plan to) because snakes scare me.

This doesn't seem to be opinion-based to me. Either snakes can get dizzy or they cannot. Am I missing something?

@Resonating - Agreed - this seems an entirely valid question.

This question appears to be off-topic because it This post appears to be off-topic because it does not suit the target audience of this site (biology researchers, academics, and students). While questions are welcome regardless of their difficulty, users should have made some effort to find an answer for themselves by, for example, reading basic biology text books or online material first.

In what context have you seen snakes get "twirled around in bags"?

@augurar I was mistaken.  I haven't seen them twirled around in bags.  I've seen the end of sacks twisted that were used to capture snakes.  I guess I just wondered what would happen if you twirled the bag around.

Probably don't do any experiments that involve shoving reptiles in a sack and swinging them around your head, as someone will surely call the ASPCA on you.
The type of dizziness that is associated with spinning around is called "vertigo". From wikipedia:

Repetitive spinning, as in familiar childhood games, can induce short-lived vertigo by disrupting the inertia of the fluid in the vestibular system

So then from a physiological perspective the question is, do snakes have fluid in their ears like people do? And a quick review of snake physiology seems to suggest that they do.
Thus, I'm going to put forward the tentative hypothesis that yes, snakes can get dizzy, but it's still probably very different from you think of as dizziness.

Dizziness is caused by a continuing rotation of the fluids in the semi-circular canals of the vestibular system that transmit rotation information to the brain. After prolonged rotation of the body, the circular canal fluids are engaged in a rotating motion. This fluid motion continues even when the body has stopped moving. The persistent fluid rotation is detected by the hair cells lining the canals that in turn send this aberrant information to the brain that interprets it as continuous motion of the body. A picture from the wiki page on the vestibular system:

All jawed vertebrates, including snakes, have this basic system of a horizontal, posterior and anterior canal filled with fluid (wiki on the vestibular system). Based on this common anatomy, hence, it is reasonable to assume snakes do get dizzy. However, as @tel rightfully indicates, dizziness is a subjective perceptual phenomenon and the assumption of snakes getting dizzy needs experimental verification. Ethically, however, this would (and should) never get through a review board as it is a curiosity-based question only.

I will try to replace the wiki citation with a more trustworthy journal reference later. The wiki cites a book I don't have access to.

Perhaps you could restrain a snake in a long tube, spin it along the long axis, and then measure its ability to strike at a target? I wonder what snakes use their vestibular system for, they are in contact with solid surfaces nearly all the time...

The vestibular system detects motion in general and sends feedback to many systems, including the oculomotor system that keeps the eyes fixed onto a potential prey. No just-out-of-curiosity-experiments on animals, please :)

There's no danger of me putting snakes in socks, don't worry.

@ChrisStronks You're right, you should submit an application to your Institutional Animal Care and Use Committee for permission to swing snakes around.

In fact all craniates have some kind of vestibular system -- hagfish have one semicircular canal and lampreys have two.

It is impossible to objectively answer this question, as there is no defined method to objectively tell if the animal is dizzy.
All that is known is the existence of vestibular system. Vestibular system helps to define ones orientation with respect to gravity.
The response of a reptile to such a stimuli would most probably be freezing as opposed to a four-footed or two-footed beings. This is due to the stabilty of posture present in reptiles which have either their entire body opposed to the ground (in the which the center of gravity touches the ground) or have their centre of gravity very very close to the ground. As the reptile is already stable there would be no movement in the form of corrective reflexes.
The bipedal or quadipedal beings on the other hand have a high center of gravity and when the vestibular system is stimulated, the dizziness will kick in some corrective reflexes which can be objectively measured.
So the answer can only be derived based on observations on humans or other creatures with quantifiable response to the stimulation of vestibular system.


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Can I drink warm/hot tap water?
I live in Canada and tap water is drinkable, as a matter of fact, the government puts a little bit of flouride in the tap water so it cleans your teeth.
My wife and mom always tells me not to drink the tap water when it's warm/hot.
Is there something in the water that's bad for the body or is this a myth? (bacterias in warm/hot water vs cold)
Added info: The water is heated in the water heater which is in the basement of my house. It is a new water heater, only a year old. My house is built 7 years ago, which for a house it's not old. I don't know about the pipes though

No, there is no reason to not do this. Lead pipes are a danger with cold water as well, and lead will only be found in old houses.

This is off topic - we can not possibly know about the quality of water coming out of your tap and its not really a biology problem

@GriffinEvo isn't it? I was thinking more towards bacterias in the water against the human body

@Huangism judging by the answers it has so far it seems more chemistry (though I can understand you posting it here) - it would be interesting to post a follow up question on the Chemistry stack exchange to see why more contaminants are released in to hot water than cold

This actually depends on how your tap water is warmed.
If you have a personal water heater, which is connected to your tap with non-corrodable pipes, then there is essentially no difference.
However, if the heating comes from a common source, then it would be very difficult to verify that the heater-tap connection does not contain any corrodable pipes. In this case, it might be better to not use the warm water for direct drinking, especially if the building is old.
A 7-year old house should not have lead pipes installed, since the Canadian Plumbing Code has banned lead pipes since 1975. As such, I believe that your warm water is safe to drink.

The National Plumbing Code allowed lead as an acceptable material for pipes until 1975 and in solder until 1986. All provinces and territories use the National Plumbing Code as a basis for their own regulations.

The water is heated in the water heater which is in the basement of my house. It is a new water heater, only a year old. My house is built 7 years ago, which for a house it's not old. I don't know about the pipes though

Edited to fit the new information

Added to the question

The reason for this worry is chemical--as water warms up, more contaminants can dissolve in the water. With water pipes specifically, people worry about lead contamination. Modern pipes are not as likely to corrode (or even be made of lead) as old pipes, so this is more of a concern in older buildings.
See this NYT article for more information.


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if the load is removed from an eccentrically contracting muscle, will it start contracting concentrically?
Geometrically speaking, a muscle contraction should exert a force between the two attachment sites. However, the actual effect of the force on the overall geometry of the body near the point of attachment may not match the direction the force pulls. For example, although the bicep contraction in principle would pull the forearm closer to the humerus, instead during eccentric lengthening the arm is actually extending due to how the rest of the system is configured. This is an example of where the direction of force exerted by the muscle doesn't coincide with the movement of the
My Question:
Can a muscle's net effect on the geometry of the system it acts on change mid-way through contraction?
For example, if my arm is eccentrically extending due to too heavy a weight in my hand and suddenly I drop the weight and now the bicep can contract. That's a simple example but I'm wondering how common it is for a the effect of a muscle's action to change midway through.

A muscle's action is one way, and not two-way as you seem to imply if I understand the question correctly. The biceps bends the arm by contracting. Upon bending the triceps relaxes. The reverse happens when the arm is flexed, i.e. the triceps contracts and biceps relaxes. Biceps and triceps are known as antagonists. Many striated (skeletal) muscles work this way, i.e., with antagonistically paired muscles.

These folks would seem to disagree: http://biology.stackexchange.com/questions/29655/do-both-ends-of-a-muscle-contract

They don't disagree with me and I not with them as far as I can see, it's kind of a huge post. Be specific in who disagrees with what and why.

It is not clear to me what you are asking.

The question in the title and the body are asking two different things.

I don't think this question is unclear. If I understand correctly, it's asking that if the load is removed from an eccentrically contracting muscle, will it start contracting concentrically.

Yes, that's exactly what I am asking. I just didn't phrase it that well. Maybe I should adjust the title

"This is an example of where the direction of force exerted by the muscle doesn't coincide with the movement of the"..... something missing here

Yes, because net force is the only thing that matters. To address @Chris's comment, in an eccentric contraction and a concentric contraction, the muscle is applying force in the same direction. However, in an eccentric contraction, the muscle is not applying enough force to actually lift the load. A biceps curl is a good model:

SportScience
In the second image, the muscle is contracting but not exerting enough force to lift the weight--the net force causes a slow lowering of the weight.

Rather than working to pull a joint in the direction of the muscle contraction, the muscle acts to decelerate the joint at the end of a movement or otherwise control the repositioning of a load. - Muscle contraction (Wikipedia)

It may be clearer with an actual force diagram:

University of Guelph
The only factor influencing which way the lever (your arm) moves is the net force. So yes, if you change the load (force F in the above image), the muscle could begin pulling your arm in the opposite direction. The muscle isn't really "changing action", it's exerting the same force, but now there's less force resisting it and the overall movement changes direction.
The question in the title and in the body are not really the same question, but the force is still exerted in the same direction as the muscle. You don't "see" movement in that direction because the forearm on which the force is being exerted is fixed at the elbow joint.

Yes.
You can confirm this by yourself easily. Put your right hand in front of your shoulder and use your left hand to push it out in front of you and downwards (keep the left upper arm in place). Resist the movement, so that your left biceps contracts eccentrically. Now snap your right hand away and you'll notice that your left hand snaps upwards (contracting concentrically).
This also works for isometric contraction (unless you're contracting at the end of the range of motion, because then the muscle cannot lengthen anymore for further concentric contraction). Do the same setup, but equalize the push forces the hands exert upon one another.
You can fonfirm this with another person exerting and releasing the force, if you're suspicious that you might be tricking yourself.


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Is the six-layer cortex model of the mammalian cortex still the most accepted model?
I've been reading a bit about the different layers of the cerebral cortex and its clear that certainly not every region of the cortex has the same number of layers. Thus, the idea that every region has six-layers is clearly false.  And I think this fact is definitely well known.
Do neuroscientists still view the six-layer model as an accurate model? If so, what aspects of it have changed over time that now incorporate the fact not every region has six layers?
Aside:
Let me clarify my question. If not every region has 6 layers, why don't we just say there is a layered cortex and not specify a number? But if we decide to do this, what distinguishes the mammalian "6-layered" cortex from non-mammalian species that have a so-called "3-layer" cortex
See The Microcircuit Concept Applied to Cortical Evolution: from Three-Layer to Six-Layer Cortex by Gordon Shepard for more.

One distinction: cerebral cortex is divvied up into isocortex (6-layered) and allocortex (3-4 layers). This (title: The Microcircuit Concept Applied to Cortical Evolution: from Three-Layer to Six-Layer Cortex) looks like a good read for the relation of three-layered and six-layered cortex.

Nice, I will check it out.

Short answer
The layers in the cortex are histologically and functionally defined. Both 3- and 6-layered cortices are found in the human brain. Hence, the different cortical layers are not models, but classifications based on empirical observations.
Background
The different cortical layers have been defined by histological staining and microscopy. The layered organization of the mammalian cortex is typically explained in textbooks by using the neocortex as an example, which has 6 layers (Fig. 1):

Fig. 1. Layered cortex. Source: What-When-How - Neuroscience.
As one can see in this picture, layer VI can be divided histologically into 2 sub-layers, namely VIa, containing mainly pyramidal cells, and VIb with mainly horizontal cells (Prieto & Weiner, 1999). Hence, a 7-layered cortex could be argued for as well - it is all kind of subjective.
The functions of the 6 layers are illustrated in Fig. 2.

Fig. 2. Functions of the 6 layers. Source: Free-Stock-Illustration.
However, the primary olfactory human cortex (the paleocortex) contains only 3 histological layers, as opposed to the 6 identified layers in the neocortex (Fig. 3). The primary olfactory cortex is part of the allocortex in man.

Fig. 3. Neocortex versus paleocortex. Source: Slideshare.
The primary olfactory cortex receives direct sensory input from mitral cells in the olfactory bulb in the outermost layer Ia. layers Ib, II, and III receive input via local and long-range intracortical connections. Sensory and intracortical inputs converge in layers II and III. The three-layered palaeocortex is different from the six-layered neocortex in sensory areas. In the neocortex, sensory and intracortical microcircuits are distributed to two different layers. Sensory inputs from the thalamus target layer 4. Layer 4 subsequently projects onto layer II/III, which distributes and receives intracortical associative fibers. Note that the olfactory system is exceptional in that it does not relay its information via the thalamus, explaining the absence of layer IV (Wiegand et al., 2011).
Reptilians and birds typically have a 3-layered cortex (Nauman et al., 2015).
References
- Nauman et al., Current Biology; 25(8): R317–R21
- Prieto & Weiner, J Comparative Neurology (1999); 404:332–58
- Wiegand et al., J Neurosci (2011); 31(34):12149 –58


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What is an apomorphy? What is a plesiomorphy? Can someone confirm these definitions?
I want to know the correct definitions for the terms apomorphy and plesiomorphy. Unfortunately, there seems to be a variety of ways to define these terms depending on the language one uses. Thus, it has been hard to determine the correct definitions. I have settled on definitions that seem correct to me and also have examples. I would like confirmation from someone if I am understanding the definitions correctly.

Apomorphy -- any character state or trait novel to a species and its descendants. An apomorphy occurs when a taxon is selected to have a particular trait. Example: within the class reptilia, the suborder serpentes (snakes) has an apomorphy because its members have no legs.
Plesiomorphy -- an evolutionary trait or character state that is homologous within a particular taxon but is not unique to members of that group and therefore cannot be used as a diagnostic character for the group. Example: within the class reptilia, legs are a plesiomorphy for its members.

My Question:
Are these definitions and examples accurate?

That sounds pretty good. On this kind of things wikipaedia is very reliable.
You can also look at this very synthetic page:
https://www.enotes.com/homework-help/distinguish-between-synapomorphy-apomorphy-304128

The link has expired.

Apomorphy can simply be defined as derived trait or character state that is distinct to a particular species or group in a phylogenetic tree down to its descendants. The example you cited is fine.
Plesiomorphy is an ancestral or primitive traits that are homologous to a certain group of organisms but not unique to the other members of the group, hence cannot be used as a diagnostic character for the group.

Please try adding citation or scientific references to your answer.


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What are the most common mechanisms motivating mammals to mate?
I read somewhere that dolphins and humans are some of the only species known to have sex for fun. I was just curious

What motivates mammals to mate? (Like what compels them to do it, not what the purpose of doing it is)
Are those motives solely aligned for reproduction or are there other benefits to mating?

There are pheromones and other hormonal signals. I don't think sex was ever about fun. IMO, fun (or the ecstatic feeling/dopamine release) is probably just a response that would increase the chance of mating (in the sense of the organism wanting to try again).

What determines when pheromones and hormones are released? Are there mating seasons where environmental factors trigger them?

There are environmental factors. It can be periodic like in the case of estrus. I have to read a bit to answer your question in detail.

Different animals are compelled by different things. If you have a specific animal in mind I'd suggest focussing the question on that. As it stands the question would require too broad an answer.

I enjoy questions on animals mating


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Is dysharmonia an actual disease or something only Oliver Sacks believes in?
In his book Musicophilia, Oliver Sacks discusses a case involving a woman named Rachel Y. who lost the ability to process harmony in a car accident. He then explains how this lead to simultanagnosia. He terms this loss of harmony "dysharmonia."
Is dysharmonia an actual medical condition or is this something Sacks just believes but has no real evidence for? My google fu didn't yield anything besides his book.  If it is legit, which regions of the brain are damaged and why are those regions in particular thought to induce dysharmonia?
Click here to read the case.

This might be better suited to [skeptics.se], I'm not sure...

Never used Skeptics so I can't really say if I agree or disagree. If it is, that's fine. I'll let others weigh in and if it is mods can migrate it.

Check out their welcome to new users post to learn about the site and how it works. You don't necessarily need to have the mods migrate it, you could just post the question there, possibly with some edits to satisfy their question criteria.