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How does it make sense to speak of heritability of qualitative traits?
As far as I understand, heritability is defined as "proportion of variation of a phenotypical trait due to genetic variation between individuals in a population".
I see the concept is being applied on quantitative that can be measured with numbers, can be compared (i.e., $\mathrm{A}$ is taller than $\mathrm{B}$ by $x$ units of height) and varies continuously to form a distribution (e.g IQ forms a Gaussian distribution).
heritability in the narrow sense ($h^2$) has a mathematical definition. For a given quantitative trait $x$ of a population that forms a distribution with certain average and certain variance $V_P$ then $$h^2 = \dfrac{V_G}{V_P}$$ where $V_G$ is the variance in population due to variance in genetics and $V_P=V_G +V_E$ where $V_E$ is variance in population due to enviromental variation (I ignored other terms in the definition of $V_P$).
IQ has mean of 100 with $h^2=0.8$. So for a person whose IQ is 105 (5 points above average), genetic contribution contributed to this $0.8*5=4$ points difference and the remaining 1 point difference is environment.
I also see how the concept can be applied on qualitative traits (by this I mean traits that don't satisfy the properties of quantitative traits) even in the absence of well-defined notion of variance $V_P$ because the trait doesn't form a distribution. For example in a population of flowers that can only be red or blue.This red/blue trait does not have a well-defined $V_P$. We can still meaningfully say that this trait has $h^2=1$ since any variation in the color (red vs blue) is due to genetic variation. We can also speak meaningfully and say a certain qualitative trait has $h^2=0$ meaning any variation in the trait is due to different environments.
On the other hand, I think qualitative traits can have either $h^2=0$ or $1$ but no value in between, since $V_P$ is not well-defined (and hence $V_G$ and $V_E$ as well are not well-defined) for them.
Consider sexual orientation for example. Based on some twin studies it was estimated that homosexuality has $h^2=0.5$. But what does that mean really? how does homosexuality vary to form a variance? A person is either homosexual or not(heterosexual, bisexual or asexual).
My question is how to apply this $h^2=0.5$ value of homosexuality in the same way I applied it above on IQ?
More generally how to meaningfully be able to interpret heritiability of other qualitative traits (mental disorders for example like schizphrenia with $h^2=0.8$) that has has no well-defined $V_P$ and whose $h^2$ value is not $1$ or $0$ but rather lies between them?

Can you give us reference for twin study on homosexuality, or even on IQ?

The concept of heritability is a concept coming from quantitative genetics. It only applies to quantitative traits. This does not mean that the trait must be continuous. Discrete traits (such as the number of eyes for example) are quantitative traits.
For boolean traits, it is common to set one outcome to 0 and the other to 1. Which outcome is set to which value does not matter as it won't affect the variance. To take your example of homosexuality, one would classify all individuals as homosexual or heterosexual and set homosexual to 0 and heterosexual to 1 (or vice-versa) and compute the heritability from there. Of course, categorising people as either heterosexual or homosexual completely misses the diversity of sexual orientation. The ability to put its tongue in the vertical position would be a better example of a boolean trait.
As far as I know, for non-boolean nominal traits, the concept of heritability is undefined.

I would imagine you could create dummy boolean variables for nominal traits, and analyze them as a boolean to get their heritability.

I think after some research I found the answer to the question. Measuring heritability of binary traits (e.g, diseases that do not conform to Mendelian rules like diabetes, mental disorders like schizophrenia and bipolar and etc) is based on the liability threshold model.
This theory assumes that every binary trait has underlying liability value that predisposes an individual for this trait. This liability varies continuously and forms a Gaussian distribution for a population. There's a threshold value of liability after which an individual becomes affected by this trait and it manifests itself (e.g, becomes schizophrenic). A person below the threshold value will be unaffected (e.g., does not have schizophrenia).
The theory posits that the source of variance $V_P$ of liability among the population is owing to genetic and environmental variances $V_G$ and $V_E$ respectively.
heritability as estimated by twin studies of monozygotic and dizygotic twins then estimates the heritability of the liability of a binary trait (e.g, homosexuality). In this case the formal definition of $h^2$ applies.

Using a threshold model based on a liability factor implies you having a liability factor and hence a quantitative trait.

Having a liability factor does not imply a quantitative trait. In fact, the threshold model is designed specifically to deal with binary traits. The underlying liability, i.e. the genetic + environmental risk, is Gaussian but the trait (e.g., having a heart attack or not, dying or not) is continuous. This is known to be the case for many diseases. A simple analogy is your risk of lung cancer increases continuously with every second you're smoking a cigarette, but the trait of having lung cancer or not is not quantitative. Binary can be considered continuous or categorical so it may semantics.


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Characters do not make the genus, but the genus gives the characters?
In his Origin of species, in the chapter entitled 'Mutual Affinities of Organic Beings: Morphology: Embryology: Rudimentary Organs', Darwin, speaking of the nature of classification systems of living organisms, writes:

Such expressions as that famous one of Linnaeus, and which we often meet with in a more or less concealed form, that the characters do not make the genus, but that the genus gives the characters, seem to imply that something more is included in our classification, than mere resemblance. I believe that something more is included; and that propinquity of descent, the only known cause of the similarity of organic beings, is the bond, hidden as it is by various degrees of modification, which is partially revealed to us by our classifications.

What does he mean by  "that the characters do not make the genus, but that the genus gives the characters"?

I cannot really know what Darwin intended to say but I guess this statement means that two distant species may look similar but they are actually evolutionarily distant.

It sounds like the intent of the phrase is to suggest that the presence of similar characteristics in two species do not imply that they are related, but that related species would be expected to share characteristics.
I.e. that similar characteristics do not imply relatedness, but relatedness implies similar characteristics. Genus -> Characteristics, rather than Characteristics -> Genus. These two possibilities are logically distinct.

@InactionPotential I think you can make an answer out of your comment.

I am doing research in marine benthic ecology and taxonomy. I think the above sentence denote that one species is originated from a single ancestor,but they have different characters for adaptation in a particular environment. that mean they have similarity in morphological characters,but genetically different.

It has been a long time since, I've read The Origin of Species, but it sounds like the intent of the phrase is to suggest that the presence of similar characteristics in two species do not imply that they are related, but that related species would be expected to share characteristics. I.e. that similar characteristics do not imply relatedness, but relatedness implies similar characteristics. Genus -> Characteristics, rather than Characteristics -> Genus. These two possibilities are logically distinct.
The assumption that Genus -> Characteristics segues into the key concept of his theory of evolution, that related species (a genus) arise from the descent with modification of an shared ancestral population. Remember that Darwin wrote long before we understand the role of DNA as the carrier of genetic information. He knew nothing about the physical substance or the mechanism of transmission that passed characteristics to offspring, but through his observations built a strong case that such a substance exists and that gradual changes in this substance occur over time in a population. He later posits that natural selection leads to the accumulation subsets of descendants with modifications that are advantageous in their environment. Thus the species within a genus could arise, their similarities the result of their shared ancestry, their differences that result of descent with modification and selection.

I'm not sure when the concept first arose but I would assume it was after Darwin had come up with the Theory of Evolution.  The concept I am talking about is "Convergent Evolution" which in essence explains "characteristics do not make the genus".
Convergent Evolution is the concept that two completely different organisms, given the same selection pressures, may, independent evolve to exhibit similar characteristics.  If one was to assume "characteristics makes the genus" one would think these two must be genetically similar when they are not.
One example I found illustrated from link is:

If you look at the external characteristics, we would say the 3 must be closely related - tail fin, dorsal fin, pectoral fins all pretty much the same.  But looking at the bones and others we know that they each evolved from very different species.  Not only are they from different genus, their divergence goes all the way up to the class level (Kingdom, Phylum, Class, Order, Genus Species).
Why then do they all look similar?  The reason is they all have to deal with the common selection pressure of water.  Living in water, moving efficiently in water to be able to catch prey and evade predators.  Separately, they all gradually evolved fins because the ones that did not, were out competed by there "better finned" peers.  The better finned shark ancestor out bred the poorer finned sharks, the better finned porpoise ancestor out bred the poorer finned porpoise.  The ichthyosaur too except it finally did not evolve sufficiently and became extinct.

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Can I explain the kinetics of a ligand looking at the ligand-protein co-structure?
Can I explain the kinetics of a ligand binding to a target protein (association and dissociation rates) by looking at the protein-ligand co-structure?
Editing my question after a few comments:
I have a set of ligands targeting the same protein. These ligands show different kinetics properties, i.e. different association and dissociation rates. Would it be possible to explain why they have a different kinetics if I had the x-ray structure of the different protein-ligand complexes? Would I be able to say that ligand A has a slower dissociation time compared to ligand B because it makes for example 3 H-bonds instead of 2? How accurate would this conclusion be? Would I need in any case MD simulations?

Theoretically, structure defines function. However, it is very complicated. When people design enzyme on paper and then synthesize it, predictions are rarely accurate.

I am not sure what you want to know exactly.

It is not generally possible to have an idea about dynamic properties from static data. I guess you can use molecular dynamics simulation based on the structural data to have some idea about kinetics (I am no expert in this area).

yes you probably need some molecular dynamics simulations. And what about if I have several co-structures with the same protein but with different ligands with different kinetic properties? Can I explain their different kinetics based on their binding mode?

Specifically what data do you have? Do you have a docked crystal structure?

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Has a restriction enzyme ever been tagged?
As part of an experiment that I am preparing, it would seem necessary for me to tag a restriction enzyme (HaeIII to be exact) with GFP. I began researching its domains to determine whether to tag the GFP to the N- or C-Terminus, but I was unable to find any previous studies where this restriction enzyme was tagged. As I got asking around, it would seem that no one has ever tagged any restriction enzyme at all.
It appears that this is only the case with restriction enzymes, because I very quickly found several articles where the HaeIII methyltransferase was tagged with GFP.
So, does anyone here know if a restriction enzyme has ever been tagged before? Is there something that I am overlooking that would inhibit the tagging of a restriction enzyme?
Thank you.

I don't see a reason why it should be generally impossible to GFP-tag a restriction enzyme. I suspect, that simply nobody has had the need to do so until now. I suggest you simply try both N- and C- terminal variants simultaneously (including a flexible linker) and test their activity.

Just out of curiosity, why are you tagging a restriction enzyme anyway?

@Thawn, I had the same thoughts. I'll just make both constructs and see which works better. I was just wondering if anyone has ever done it before. Like I said, I found several articles that tagged GFP to the N-terminus of HaeIII methyltransferase, so I was able to make one construct that worked well. I was hoping to do the same with the restriction enzyme, but it's alright, it won't be too much of a hassle to make both.

@MattDMo, just to make sure the cells are transformed properly. Of course, I need to introduce the methyltransferase before the restriction enzyme, so I need both to be tagged with a fluorescent protein to select whichever cells express both. Additionally, we will be observing the concentrations of both the methyltransferase and the restriction enzyme in different parts of the cell, so fluorescent tags are a win win.


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What causes DNA gel electrophoresis bands to swirl?
I recently ran a gel on a new system and came out with something I've never seen before.
I loaded the first three wells with a ladder in the first and an uncut plasmid in the next two. There are no bands, just swirls all over the gel.
It's a 10 cm gel, 0.7%, and I ran it at 80 v for 1.5 hours. Part of the new system is the buffer, lithium borate,  which I've never worked with before.
Any ideas as to why this is happening?

Edit: thank you for your answers. I followed the regular protocol closely and used the same bottle of buffer for both making and running the gel. I did this twice and got similar results both times.
I had the power supply tested and discovered that the voltage may have been fluctuating between 77 and 83 volts during the run. Could this have caused it?

Are you sure you used the right buffers or prepared the correct (dilution, concentration)?

I didn't prepare the plasmid, but I know the ladder is correct. Yes, lithium borate was all it called for, and I diluted it as the instructions said.

What length of DNA fragments are you expecting to see? What is the power (Watts) or current (Amperes) in your region?

It can possibly be one of two problems:

Either the gel poorly prepared, in that it was cooled for too long, resulting in parts that solidified before pouring it into the mold.
This I have seen before, the pH of the electrophoresis buffer and the buffer used to make the gel was not the same. It has to be the same throughout the gel to ensure that the DNA moves evenly to the positive electrode.

Do you perhaps have a gel image of the bio-transilluminator? It could provide more insight.
Hope this helps

Because the question says that a new lithium borate buffer was used, we should also make sure that the gel was made up in LB buffer with the same final concentration as the running buffer.

Electric potential (voltage) should apply equally across the gel. This should not cause DNA to run unevenly. In gel electrophoresis, the electric forces affect all (negatively charged) DNA molecules and the flow in the same direction.
What causes differences in their flow is the resistance to diffusion or flow of DNA molecules by the gel. Larger molecules are more likely to collide with the polymer matrix and will therefore move through the gel more slowly. If you have DNA prepared in varying sizes, they should form well-defined bands. There are several parameters that you can optimise, depending on the sizes of the bands that you want to distinguish:

concentration of the gel: a denser gel will slow DNA molecules more and is useful to prevent smaller molecules running off the end of the gel
time or voltage: smaller molecules running on a denser gel will take longer to separate so you may need to adjust the running time or voltage accordingly.

It is possible that the gel or input DNA has not been prepared correctly. A gel needs to be mixed and poured while it is still liquid (before it cools) to ensure that the polymer forms evenly. However, the absence of a positive control or DNA ladder even in the wells shows that the DNA is not present in this gel at all: It has run off either end of the gel. Therefore either:

the DNA ran through the entire gel: it was run for too long or the polymer was not a high enough concentrate. Prepare a gel with a higher concentration or run it for less time.
the DNA ran the wrong way: please check that the electrodes were connected in the correct orientation (negative by the wells and positive at the other end). These should be colour-coded (usually red and black).

It seems that the gel was not prepared properly and does not have a homogenous  texture as it should, which then affects its pores and prevents the sample from traveling through (hence be absence of bands).

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Effects of tobacco smoke on plant health
We know that tobacco smoking is associated with a wide range of negative health effects in humans, which holds for active smoking, second hand smoking and probably even third hand smoking.
What are the effects of second hand smoke exposure on plants?
For example the german wikipedia page on nitrogen dioxide claims that exposure to nitrogen dioxide leads to reduced yield in apple and pear trees, white birch, barley and salad. It is also stated that first physical signs of alterations can be detected with one time exposure at levels as low as 3 ppb for less than one hour. However I cannot access the source because it is a book.
Bonus: How does cannabis smoke differ from tobacco smoke with respect to plant health?

Plants have varied reactions to smoke of different kinds, either to grow differently or to fall ill. this page says that tomatoes and peppermint contain nicotine and nicotine is used to treat plants, so the nicotine can't be that detrimental to them, but the other many smoke compounds can make them ill, but only in very high levels of smoke. http://www.takepart.com/article/2015/04/15/nicotine-plants-second-hand-smoke

https://www.vidarholen.net/contents/junk/nicotine.html

@CuriousIndeed Your question seems to be asking about the gaseous components of cigarette smoke and their effects on plants, but, considering a house plant exposed to concentrated cigarette smoke, I think the particulate matter in secondhand smoke is likely more immediately harmful to plant health. Consider the effect that any particulate matter will have:  residue or tar that settles on the leaves will both inhibit photosynthesis and clog stomata, reducing the efficiency of gas exchange.

Results from the peppermint plants study show that all the peppermint plants contain minor amounts of nicotine before treatment, but the experiments revealed that the plants also incorporate nicotine considerably from the soil as well as from tobacco smoke. These findings demonstrate for the first time that the reported occurrence of nicotine indeed may originate from tobacco. The incorporated nicotine was subsequently metabolized by the plants.
Effects:
Apart from the nutritional aspects, the results on nicotine uptake may also affect basic plant biology, because they demonstrate that alkaloids can be transferred from one plant, after its death, to another plant species.

The plants, said Dirk Selmar, the study’s lead author, appeared, at
  first, to be unchanged. “As far as we have seen, there have been no
  negative effects on the growth,” he wrote in an email. Yet, tests for
  nicotine, conducted at various stages in the plants’ life, found that
  they had “tremendously elevated nicotine levels,” Selmar said.

References:
https://link.springer.com/content/pdf/10.1007%2Fs13593-015-0298-x.pdf
http://www.takepart.com/article/2015/04/15/nicotine-plants-second-hand-smoke
https://www.ehow.com/info_7987584_effect-cigarette-smoke-house-plants.html
some anecdotal answers (to be taken with a grain of salt) on your 2nd bonus question: https://www.rollitup.org/t/is-smoking-weed-around-a-growing-plant-bad.182179/

The answer focused too much on nicotine an ignores the rest..considering smoking, the health hazards of nicotine are negligible compared to the other constitutes of smoke..e.g Polyzyclic aromatic hydrocarbons (PAR), Nitrosamines, Aldeshydes, Arsenic, Nickel, Cadmium..Thus I'm not particularly happy with the answer however because its the only one so far the bounty goes to you if no one else answers...

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Why are certain aneuploidies more common?
Certain aneuploidies such as trisomy-21 (Downs syndrome), trisomy-18 (Edward syndrome), Turner syndrome (XO) etc are more common than others.
I had a vague thought that it is related to chromosome size: the chromosomes which are towards the extremes of size will be affected in an improper segregation event.
Chromosomes 13, 18, 21 and Y are among the smaller chromosomes in humans, and are commonly associated with aneuploidy (but 22 which is also small is not affected). The larger ones are unaffected (X is an anomaly).
I thought that longer ones should be more affected because more cohesin bridges have to be broken, but it seems that is not the case.

Is there any association of chromosome size and frequency of aneuploidy?

EDIT

If the difference is because of differential lethalities then does the length correlation still hold true? Smaller the chromosomes, lesser the genes and hence lesser would be the effect of extra copy on the cellular biochemical networks.

re: lethalities, I'd say it still depends on the genes and the dosage (extra or missing copy). Not all gene and dosage combinations are equally essential/lethal.  I don't think average "biochemical activity" is going to be useful.  What would be interesting is if someone had measured the fitness of specific gene deletions/duplications.  Also, are genes "essential" for the completion of embryonic development equally distributed among the chromosomes? Unequal distribution would obscure any length correlation.

I thought it was just that the other aneuploidies were fatal. They occur as often but miscarry earlier in the term therefore you don't see them. So basically there are only three that are viable 21,18 and the sex chromosome with a number of combinations: XO, XXY, XYY and XXX.
More information here:
http://www.nature.com/scitable/topicpage/chromosomal-abnormalities-aneuploidies-290
With regard to the frequency for each chromosome:
For males (i.e., sperm; source):

The mean disomy frequency is 0.15% for each of the autosomes and 0.26%
  for the sex chromosomes. Most chromosomes analyzed have an equal
  distribution of disomy with the exception of chromosomes 14, 21, 22
  and the sex chromosomes, which display significantly higher disomy
  frequencies.

Size of the chromosome is frequently reported as a factor in the frequency.
For females (i.e., eggs; source):

The data is obviously harder to attain. One study from IVF treatments
  show differences in the frequency of different chromosomes at
  different stages in meiosis.

there is some statistics in the link that you provided.. it doesn't actually say that other aneuploidies were reported in stillbirths or spontaneous abortions

this is still quite a likely explanation - there is very little or no data for pregnancies that terminate before the foetus is smaller than a pea.  Almost all extra chromosomes for instance come from sperms and ova formation and they may simply not compete for fertilization.

This is the answer. They all happen with similar frequency, however most are lethal. The less lethal the more common, hence trisomy 21 or Downs being fairly common; however most trisomy 21 foetuses do die before getting to the second trimester.

@AndroidPenguin.. The two papers in the answer suggest that the probabilities are not uniform..
It seems so that the aneuploidy prone chromosomes are relatively smaller..

Trisomy 13 also doesn't always lead to miscarriage / stillbirth


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Mathematical models of gene editing using CRISPR
One confusing thing I have found when reading articles about possible CRISPR based gene therapy treatments in humans is that there is vey little discussion about what percentage of your cells will actually have their DNA edited, the rate at which the editing takes place, and how to quantitatively estimate how many edited cells would be needed to "cure" a specific disease.  (It's not so confusing how edits could be made to a small embryo, what's confusing is what is actually possible for grown patients)
Can someone provide a few basic facts that calibrate what is currently possible and what is needed.
For instance, is it currently possible to edit more than 50% of the cells of an adult  nematode (or more complicated organism) using CRISPR?  How long does it take (minutes, hours, days, weeks).
For the current list of human diseases that are possible targets for gene therapy, how many cells need to be edited for one of those diseases to be "cured" (hundreds, thousands, billions, etc)
But in general is there a name for the mathematical models that try to predict how many cells will end up being edited, how those edited cells will multiply and replace diseased tissue,  and how many edits are needed to generate a concentration of missing protein for a treatment to be successful?
cheers!

There are several probabilities that you need to take into account.

The efficiency of DNA transformation of CRISPR encoding DNA and
target DNA into the host cell (varies between cell types and
transformation agent).
Efficiency of cutting by CRISPR at target site. (varies by DNA compaction - which varies by site and cell type, and guide RNA)
Number and cutting efficiency at off targeting sites.
Efficiency of degradation of free DNA by cellular antiviral systems
(varies between cell types)
Efficiency of DNA repair machinery to fix dsDNA breaks. (thus removing CRISPR cut sites. Varies by cell type)
Efficiency of recombination machinery (homologous
recombination and microhomologous recombination) (which is used to
insert new DNA sequence).
Probability of NHEJ damaging target site. (which can delete the CRISPR cut site)

Too many things to count.
However to give you some idea. The efficiency of CRISPR making a correctly targeted knock in for DT40 (chicken lymphoma) is  ~1% of all resistant colonies (ie colonies that have taken up the DNA seq used to make the knock in), when lipofectamine reagents are use. Lipofectamin trasfromation efficiency varies depending on the size of the transformed DNA, host cell type and plating density. So for a 5kb construct that transformation efficiency varies between 30%-50%.
CRISPR is nearly 100% efficient in yeast cells, ie all knock in cells are correctly targeted. This is likely due to the high levels of HR in yeast


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Impact of Alan Turing's approach to morphogenesis
Shortly before his untimely passing, the computing pioneer Alan Turing published his most cited paper The Chemical Basis of Morphogenesis (1952).
The central question for Turing was: how does a spherically symmetric embryo develop into a non-spherically symmetric organism under the action of symmetry-preserving chemical diffusion of morphogens (as Turing calls them, an abstract term for arbitrary molecules relevant to development)? The insight that Turing made is that very small stochastic fluctuations in the chemical distribution can be amplified by diffusion to produce stable (i.e. not time varying except slow increases in intensity; although also potentially time-varying with 3 or more morphogens) patterns that break the spherical symmetry.
The theory is beautifully simple and abstract, and produces very important qualitative results (and also quantitative results through computer simulation, which unfortunately Turing did not get to fully explore). However, even in the definition Turing discusses some potential limitations such as ignoring mechanical factors, and the inability to explain preferences in handedness. The particular models he considers -- a cycle of discrete cells and a circular tissue -- do not seem particularly relevant. As far as I understand, the key feature is his observation of symmetry breaking through small stochastic noise and instability.
What was the most important contribution of Turing's paper to developmental biology? Is his approach still used, or has the field moved on to other models? If his approach is used, how was the handedness problem resolved?

our current understanding of plant morphogenesis seems close to this idea (and there's no problem of handedness). See my answer here.

You may want to look into work on how the first HOX genes trigger differentiation.

This is a very interesting question. Many people have researched this topic, and many still are. But regardless, I had never heard of Alan Turing's contributions, so thank you!
First of all, I cannot actually find who first coined the term morphogen. Though people had hypothesized that chemicals could play a critical role in development through much of the 20th century, I cannot actually find the first person to use morphogen. But the most important paper really came from a guy named Lewis Wolpert, who came up with the model of a gradient of morphogens leading to differential cell fates. The idea being that if some area of an embryo produces a morphogen at a very high concentration, then as you move away from that area, the concentration goes down. So if this morphogen is required at or above a certain threshold for activity, then only those cells with that concentration will have a certain cell fate, while at lower concentrations, the cells can become something different.
But this does not really answer your question. You are asking how a single cell, which is spherically symmetrical, can determine a particular axis. Though most organisms do this is in slightly different ways, the most common feature is that sperm entry point breaks the symmetry. The best way to explain this is to show you a diagram of Xenopus (frog) eggs.

Image from: http://studentreader.com/nieuwkoop-center/
The Xenopus egg, first of all, is inherently not spherically symmetrical. There is a black animal pole, and a white vegetal pole. The sperm can only enter a marrow region of the egg about 30˚ north of the animal/vegetal line. Upon fertilization, an event occurs where the pigmented areas turn toward the sperm entry point, leaving a gray crescent. Nearby the gray crescent, in the vegetal pole, a structure called the organiser develops. This organiser creates many of the morphogens that then pattern the rest of the embryo.
Researchers have studied this a lot in many different organisms, but a few things really remain constant: eggs are not exactly spherically symmetrical, and the sperm entry point provides asymmetry.

Thank you for the answer. Any thoughts on the issue of handed-ness? Because a sperm entry point would still leave the egg achiral.

May I ask you how can I find more information about this topic?

I would think this is very much still "used." 60 years later, we finally have the first experimental support for it:
In this blog article about this journal piece the authors studied the ridges that form on the roof of mouse mouths. They manipulated the signaling molecules that induce their formation and observed changes in line with Turing's theory. Of course, this doesn't preclude other mechanisms from occurring, but supports that of Turing.

Welcome to Bio.SE!  When linking to articles you should summarise the content in your answer in case the article isn't available in the future.

Thanks. It's not my field of biology (so I'm not an expert) but the authors studied the ridges that form on the roof of mouse mouths. They manipulated the signaling molecules that induce their formation and observed changes in line with Turing's theory. Of course, this doesn't preclude other mechanisms from occurring, but supports that of Turing.

While Turing did not include mechanical effects on the differentiaton/patterning in his seminal work, other researchers have expanded on that idea. These do not necessarily change the premises that led Turing to his conclusions, nor make his contribution less relevant today. Adding a dynamic component or mechanics to the substrate in which the diffusion-driven instabilities happen can simply yield more complicated solutions.
Clear examples of this interaction between the reaction-diffusion mechanism and the mechanics or boundary conditions can be found in several places: the formation of palatal ridges in the roof of the mouth in animals, teeth formation across species, bacterial colonies both natural and synthetic and even the issue of boundary conditions has been explored in animal coating patterns.
In the case of handedness, I think this also a question of symmetry breaking, such as with Turing patterns, but happening at different scale. Skeletal primordia are thought to be formed through a Turing-type mechanism. The "issue" with handedness would be that in Turing's formulation all fingers would be equal, but superimposing the patterning already created by the Spemman organizer which polarizes the hand, makes the fingers distinct. So, in essence, a prior symmetry breaking event (which could be caused by a different lateral inhibition process or diffusion-driven instability acting a larger scale) can modulate a downstream developmental process.

Turing patterning has had a large impact in systems biology understandings of morphogenesis. The general idea is that Turing mechanisms can be coupled with other mechanisms to build robust patterning methods.
So while not too many people would say that an entire mechanism is ONLY due to Turing patterning, a lot of regulatory networks can be understand as having the components allow for a Turing patterning mechanism (under certain parameter constraints), and thus they can understand "why" certain interactions exist and predict the behavior of perturbations. Usually there are multiple mechanisms involved in a patterning system so doing some knockouts will not get rid of patterning, but by disturbing the Turing mechanism you will many times see a decrease in the robustness of the patterning.
A lot of systems biology has focused on robustness: what kinds of network motifs allow for robust switch-like behavior? etc. The Turing mechanism is a network motif which allows for robust spatial behaviors.


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Does the human body need fat for other uses than to cover essential acid requirements?
Linoleic acid and alpha linoleic acid are the two essential fatty acids, from which other fatty acids can be synthesized internally. See for example here: Essential fatty acids. Given that the requirements of these fatty acids are met, what other function do dietary fats provide which are unique to this macronutrient?
To clarify I'm asking about functions that must be provided by dietary fatty acids that cannot be provided by fatty acids synthesized from acetyl CoA from other dietary sources.

Surely by the definition of the word, essential, there can be none regarded as unique.

I have heard about fat being necessary (?) for the absorption of certain vitamines

But this is none-specific, and fatty acids can be synthesised from acetylCoA generated by the oxidation of carbohydrates etc.

You really need to clarify your question if you expect anyone to answer it with or without a bounty. Are you asking whether there are functions that must be provided by dietary fatty acids that cannot be provided by fatty acids synthesized from acetyl CoA from other dietary sources? (Incidentally, do you know what acetyl CoA is?) we can help you if we know what you are asking. If you don't know, you shouldn't be asking.

Fats can serve non-metabolic functions as well, for example structure and support around organs, padding or insulation.


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How do similar looking carps, but belonging to different species, differ from each other?
There are some carps which differ in colour, or has some easily noticeable morphological differences. But apart from that, most of the carps look "almost" the same. How does one differentiate one species from another?
Citing behavioral differences, differences in morphological characteristics (maybe subtle) and the likes which addresses this problem would be helpful. The characteristics with broader scope would be better.

This might actually be a more complicated question. The phylogeny for carps and other members of the Cyprinidae family have not been well resolved. Also, "Carp" belong to numerous genera including Carassius, Cirrhinus,  Ctenopharyngodon, Cyprinus, Hypophthalmichthy, Mylopharyngodon, etc. so they're not all actually related (below the family level). I would assume you could find a fairly easy way to differentiate these genera from each other. But...Given all this, are you seeking to differentiate all carp or just Cyprinus species?

I framed this question mainly keeping Cyprinus species in mind.

There isn't actually. Every species of Cyprinidae has its own distinct characters and sometimes for the ones that are too similar, like Labeo rohita, Labeo bata and Mrigel one can only distinguish them after regular examination. There's no set rules/guidelines but ones you develop an eye you know what ways they are different. Yet knowing the fin formula can help. But learning them all for so many species and remembering them is absolutely beyond human abilities.

If you are interested in some specific species. I think this can be answered.

Let's consider a school of fresh water carps which is readily available. I would like to know of some specific characteristics which are more effective than others when it comes to distinguishing between them i.e. if most of them can be told apart using a trait then it is significant. Developing an eye is just an observation, deducing inferences and standardizing them in a way, so that they can be used to understand subtleties, is what science should be all about.

Do you have a region of interest? As above commenters have mentioned, Cyprinid ID's are really hard. My general algorithm for fish ID typically involves a gestalt feel for family, and, depending on my familiarity with local members of the group, either referencing my internal catalog of species present (incorporating local abundance) or finding a field guide or key. But identifying species from afar? That's not going to be easy, esp. Not without some baseline familiarity with morphology and behavior.

I've edited the question.


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Can a portion of human skin cells be modified in some way to generate light?
I have come across some species of living organisms who are able to emit light at whim. Can that ability be incorporated into a portion of human skin (a specialized tissue)?

Genetic modification of any cell to overexpress a fluorescent protein will not 'generate' light, per-say: fluorophores absorb particular wavelengths and re-emit in less energetic wavelengths, dispersing some energy as heat. (Perhaps there is some semantic choice here, but there is no transduction to light from another form of energy.)
Bio-luminescence, on the other hand, can be considered generation of light. There are proteins commonly found in marine animals (which skin cells could be engineered to express) which convert chemical energy to emit light.
With regards to generation of electricity, I think this gets a bit interesting...
The electron transport chain (ETC) in the inner membranes of our mitochondria carry out a series of redox reactions which serve to oxidise food metabolites (thus freeing electrons). In order to maintain the free energy gradient that favours this directional cascade of reactions, electrons and protons are finally accepted by a continual supply of oxygen atoms, producing water. (You may be aware that aerobic respiration consumes oxygen and produces water.) In nature (in aerobes), oxygen acts as the final electron acceptor, or an electron sink.
Bacteria also have electron transport chains for their respiration (mitochondria were once bacteria, after all), and people have wondered whether these redox reactions could be used to produce a current instead of reducing oxygen to water as the final step. We normally use current in the form of electrons moving along a metal wire.
This is the idea of a "microbial fuel cell". Just like a battery produces a current from a chemical redox reaction, the ETC might be recruited in to producing a current from its chemical redox reactions.
I think this is a very difficult piece of bioengineering, since the ETC is integrated in to respiration and needs to be de- & re-coupled to some kind of inorganic mediator and ultimately a circuit; and also because the ETC produces very low voltages, I think.
There is a nice wikipedia page on microbial fuel cells that you might find interesting, if you think this is relevant to your question.

I understand your point from this answer all right.But please add a para explicitly stating "Is it possible for humans to generate electricity and light from skin tissues and natural energy, similar to that of those animals by any means, using current state-of-the-art knowledge about Biology?"

The part about using luciferase to produce light is only a partial answer. Expressing the luciferase is the easy part. Producing enough luciferin to actually make visible light would be very hard, since the luciferin biosynthesis pathway isn't fully understood yet, and is pretty energy intensive. Trying to regulate the lighting would be very hard, so even if you could get sufficient luciferin production, don't plan on turning it off, or flashing it, or anything complex.

a.aniq, your question needs a lot of work, in my opinion; it's not clear what you really want to know. Are you really asking: "Can I have skin like a fire-fly, or an electric-eel?". The answer is no, for the reason Kendall gives - these organisms have quite extensive physiological adaptations in order to luminesce or produce electric shocks, tissues very unlike human skin. My answer was instead about transducing energy and the idea of a microbial fuel cell, as I wasn't sure what you were asking, and since these small steps are more realistic.

Sort of problematic is the "at a whim" part. Just a brief search to wikipedia on electric organs, here, we see that the electrocytes are controlled by a nucleus of pacemaker neurons. This is all unique to electric fish, and you'd have to express that whole system in a human being to elicit the same effect. You'd also have to consider how your organism is engineered to withstand the actual electric shock emitted by the electric organ.
We can already tag molecules with luciferase to emit light as a mode of bio-luminescence. To get an appreciable reaction, or rather an appreciably bright light, I can imagine we'd need a ridiculous amount of luciferin in vivo, though. We also get stuck at the part about it being on a whim. The control mechanisms/structures for the light-emitting reaction simply aren't present.

As noted above, the problem with bioluminescence is not in producing luciferase in vivo - I have a mammalian plasmid in my lab that can do that. The issue is producing and trafficking luciferin (luciferase's substrate).

Yes, the fibroblasts of skin cells can quite easily be Transfected with a fluorescent gene such as GFP. Fluorescence doesn't actually produce light, the fluorescent molecule just changes the wavelength of light shining on it.
Fibroblasts can also be made luminescent (light producing) by acquiring  genes for firefly luciferase or aquoren from jellyfish. This is a little more tricky since luminescent molecules typically require cofactors such as ATP, Calcium or other proteins.
Both methodologies are readily performed in vitro.

And what about electricity?

That's a much more complicated physiology based on  tissue arrangement. Although I'm not 100% on this I think introducing a phenotype like that into a transgenic animals at this point is still science-fiction

I don't know if what you are saying is possible or not. I'll wait for somebody to comment on this answer, before thinking on it.


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Can genetically engineering the DNA of a human zygote, make it a twin of another human in entirety?
My question is simple. If I wanted to make an exact twin of an individual, will genetically engineering the DNA of the zygote to match with the individual suffice?

Can you please expand and better explain your questions? At the moment they are often hard to understand.

Edited. Is there any more doubts?

Is it possible, from any cell of an individual? (speaking theoretically) Then are you saying it won't be an identical twin if just the DNA contained within the cell was identical? If so, which traits would differ specifically?

No, playing with the zygote will not do it. It needs to be the nuclear material of the individual you want to clone inserted into a donor egg after it's nuclear material is removed. When one starts dividing, the embryo can be transplanted into a surrogate mother. Read about Dolly the sheep.

Yeah, are you asking about strange cloning variation?

The answer is no, and it is because sequence of the genome is not all information that is required for gene expression and development, there are also epigenetic factors.
A lot of patterns of epigenetic marks, such as most of DNA methylation and some histone modification patterns, are set during parental germline development, and these marks are transmitted to the zygote by gamets (except paternal histones, those are not transmitted because they are temporarily substituted for protamines in sperm for greater compaction). Therefore, the naked DNA inserted into zygotic nucleus will not have access to the machinery that established epigenome during development of the gametes.
And even though nucleosomes may assemble in the zygote on the naked DNA and histones may acquire post-translational modifications, and cytosines may get methylated, but most definitely the zygote is going to have no means to correctly set distinct patterns of epigenetic marks at alleles that are normally imprinted, i.e. those alleles whose epigenetic state depends on the gender of the organism that produced the gamete.
Therefore, the twin created in this way will have the same DNA sequence, but it will be a  subject to disorders associated with imprinting

If the epigenetic state of the alleles depends only on the gender of the organism producing the gamete, then it won't matter, or rather should it?

Majority of genes in gametes have similar epigenetic marks irrespective of the gender that produced them. However a small number of genes are modified differently in female gametogenesis in comparison to the male gametogenesis (i.e. the sexually imprinted genes). And I don't think that there is a way for such differential pattern to be established correctly in the zygote in your hypothetical experiment.

From your answer and comment, I am assuming that the epigenetic marks are the same for a gender (of a particular species, just for safety an extra assumption). And, for a cell to avoid disorders associated with imprinting it should have the same epigenetic markers, before it can undergo successful cloning process. So, if an in-vitro fertilization is done of two healthy gametes each from a male and a female humans. It should have the same epigenetic markers as any other fertilized human egg. Because here, as you say, in case of epigenetic markers, only the gender matters, not the individual.

DNA duplication (imagine, we PCR it) will net be enough. First of all, you also will have to duplicate and pack mitochondrial DNA into the cell. next, as been mentioned, epigenetic modification (methylation) and histone packing are crucial for proper development.
And these markers should essentially replicate state of the host cell.
My best guess is that all-synthetic replication atomic-resolution of zygote will be enough, but not just bare DNA code (750MB of information)

Mitochondrial DNA is also a DNA. Apart from that, the mother could be kept the same, if that becomes a major hurdle.


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Finding Rna-Seq dataset for C. Elegans (Caenorhabditis elegans)
I want to find a RNA-Seq dataset for C. Elegans to use it in RNA-Skim software. For those who does not know RNA-Skim let me explain it a bit. It is a tool which is used to quantify RNA-Seq data using sig-mers(special kind of k-mer). Firstly RNA-Skim clusters protein coding genes and find substrings that are special to cluster of that gene. After this preperation stage, it takes an RNA-seq data as an input and tries to quantify transcripts in it. So, I need to find a dataset in fasta file format for this step. Since I'm pretty new in this area, is there anyone who could provide me information about how could I find it?
Complete information about RNA-skim can be found here : RNA_SKIM
EDIT
The type of a dataset I'm looking for is something like that: http://codepad.org/n3ioT3nj .This dataset is for mouse, I need to find such a dataset for C. elegans as I said before

Try any of these: www.wormbase.org, www.modencode.org, data.modencode.org, or ftp.modencode.org. Also, they won't be fasta files for NGS data they will either be fastq format or SAM/BAM format

Go to NCBI-SRA or GEO and search for RNAseq data. Set organism as C. elegans and optionally any other keywords that you are interested in. This search term will give you all RNAseq data in SRA from C. elegans: ("caenorhabditis elegans"[Organism]) AND "strategy rna seq"[Properties]

You will generally find files in fastq format which you can easily convert to fasta. There are many tools that do that; you can also do it with your own script. Following awk command will do the job:
awk 'NR%4==1{print ">"substr($0,2)} NR%4==2{print}' data.fastq > data.fasta
Apart from SRA you can also look at ENA (European Nucleotide Archive) and DDBJ

,thanks for information.As you said, I searched on the NCBI-GEO and when I typed C Elegans RNA-seq, I saw 65 results in this link : http://www.ncbi.nlm.nih.gov/gds/?term=C%20elegans%20rna-seq  , from them I clicked the first one however I could not locate the fasta or fastq file you mentioned earlier. Could you help me ? As it is my first time interacting with such databases I have some difficulties.

SRA is the right place for sequencing data. GEO will have all kinds of expression data. In SRA you just have to click on a particular run and find the experiment ID (Starts with SRX). Paste it in this page and download the data.

@stackunderflow In the GEO page when you click on one of the experiments you will get a SRA link at the end of the page. You can click on that (ftp link) and download your file. But these will be in .sra format which you have to convert to .fastq. .sra files are smaller and will download faster and you can use fastq-dump of sra-toolkit to convert sra to fastq. Or else paste the experiment ID in the second link that I mentioned in the previous comment.

thanks a lot, I started downloading a fasta file from the link you provided. I have one more question, As you know, I've tried to find a rna-seq data and I have already shared rna-seq data for the mouse specy in my question (http://codepad.org/n3ioT3nj ) Now, I tried to figure out what do these numbers in that file explain to me. I mean how could I understand expression level of something by looking that file. For example I could not understand nothing from the line 1:4807788-4897909W:ENSMUST00000150971:8:789:94:209:S/1. How could I learn it?

And also, mouse rna-seq data that I share link of it to you is just 160.2 kb but the file I'm currently downloading already exceed the 620 Mb. Do I download wrong thing ?

@stackunderflow ENSMUST00000150971 is an Ensembl transcript id. It is your fasta header and I am not sure how you got it like that. RNAseq fasta headers are not like that. They usually have information about the platform and run details. Is this fasta file generated after mapping? Then in that case the header denotes the alignment details i.e the transcript id, location -> 1:4807788-4897909 for genomic location i.e in chromosome 1 from x to y and then the location in the transcript. I am not sure why 4 numbers in the transcript location. S/1 and S/2 are for mate pairs.

this file was not generated by me. I got it from the following web page : https://github.com/zzj/RNASkim/blob/master/src/test_data/fa_reader_test.fasta.1 .I guess they put it as an example for mouse specy but they do not provide any information about this file. Unfortunatelly I could not run RNA-SKIM software for mouse species since it requires 48GB memory at least. So I decided to use C. elegans and in order to use it I have to generate such RNA-seq data for the C. elegans species. Since I have limited characters here, I summarized it here: http://codepad.org/NSIw0Oq2

in the first step, I made a typo, it is : they clusters genes according to sequence similarities*

do you have any idea ? Actually, If I know what are these 2 seperate two files, I think I'll figure out the rest. These are 2 seperate RNA-seq dataset or part of a single rna-seq data ? If the latter one correct, why they are seperated into 2 files. I could not understand.

Actually, I got it right know.  I think they use ,as you mentioned earlier, sra-toolkit to generate such 2 files. Since I'm still downloading sra file I have not seen the result of sra-toolkit yet, but I've read that fastq-dump executable of sra-toolkit produces 2 fastq files with "1" and "2" read suffices. These suffices are also consist in the name of files that comes with RNA-skim software

@stackunderflow Paired end sequencing data has 2 files and reads in one file will have the suffix "1" in the header while the other will have "2".

I have another question related with my main question. As you can see from the link http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE60731 I have found a RNA-seq data. In the sample sections, it is written that P7 Cerebellum DNase-seq Rep 1. There are also Rep2 and Rep3. What are these Rep extensions used for ? I could not find any information on the web. Could you please explain it?

@stackunderflow These are replicates of the samples. Sometimes same sample is loaded twice and you have two runs for the same sample. This is not the case here. Rep1 is one sample and Rep2 is another sample for the same experimental protocol. If you read the paper corresponding to the data you will understand. See the study summary (click more).

thanks for the information, according to your answer, there are two possible cases, in first case I'll have two runs for the same sample. Is there any relationship with number of runs that I have and type of reads (Paired-end or single)?  Furthermore how could I know whether these "rep"s are two runs for the sample sample or they are different samples ?

@stackunderflow two runs of the same sample will be under the same experiment SRX but the biological replicates will usually have a different experiment ID. These things are usually described in the summary

I think I have found 4 runs of the same sample in this link : http://www.ncbi.nlm.nih.gov/sra/SRX145660 ( 4 of them have same SRX ID as you said). But all of them have different sizes. I guess these 4 file are not identical right? If I make RNA-seq quantification experiment on these SRR files seperately, will I get same results from all of them ?

@stackunderflow they are same samples loaded multiple times. You can analyse all of them. The profile would be the same.

Okey, if they're same I do not need to do it, but I wonder why they have different file sizes while they are same samples ?

@stackunderflow because the loaded amount may be different.


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blastn: What substitution matrix is used?
I'm currently working aligning sequences, and I need to compute similarity between pairs of DNA 'words' of a particular length.
For amino acids I am able to use the substitution matrices in Biopython (Bio.SubsMat.MatrixInfo).
However, I haven't found anything similar for DNA, so I read up and found that most systems use a match/mismatch scoring system where each nucleotide match and mismatch is scored and then the scores are summed. This works fine as long as I am only dealing with A, G, C, and T, but I run into problems when I get a sequence containing N or M and the like (meaning nucleotide unknown).
Is there a standard way to handle the situation with unknowns? That is, how do I score A versus N or M versus N?
Thanks in advance.

BLASTN does not use a substitution matrix. There are scores for match, mismatch and gaps which you can also define.
There is no feature available as of now to allow scoring of matches against unknowns. They are considered mismatches (as shown below). If these unknown are in the middle of a HSP, then you can probably re-score the HSP according to your scheme using a python script. If the N stretch is disrupting the HSP, then you can try relaxing the mismatch penalties and reduce word size (basically reduce stringency). I can't think of any other solution.
Query  1    CAGCGTCCANNTCCCGAGGTGCCGGGATTGCAGACGGAGTCTGGTTCACTCAGTGCTCAA  60
            |||||||||  |||||||||||||||||||||||||||||||||||||||||||||||||
Sbjct  8    CAGCGTCCACCTCCCGAGGTGCCGGGATTGCAGACGGAGTCTGGTTCACTCAGTGCTCAA  67

Query  61   TGGTGCCCAGGCTGGAGTGCAGTGGCGTGATCTCGGCTCGCTACANNCTCCACCTCCCAG  120
            |||||||||||||||||||||||||||||||||||||||||||||  |||||||||||||
Sbjct  68   TGGTGCCCAGGCTGGAGTGCAGTGGCGTGATCTCGGCTCGCTACAACCTCCACCTCCCAG  127

Query  121  CCGCCTGCCCTGGCCTCCCAAAGTGCCGAGATTGCAGCCTCTGCCCAGCCGCCACCCC  178
            ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Sbjct  128  CCGCCTGCCCTGGCCTCCCAAAGTGCCGAGATTGCAGCCTCTGCCCAGCCGCCACCCC  18

I understand that, I'm using the term matrix because I am handling it with one programatically, but I'm generating it with said match and mismatch values. I'll modify the question to be more specific on what I want.

@ChrisChambers Edited. But I'm afraid what you are looking for is not possible without additional scripting.

"They are considered mismatches (as shown below)" This is what I wanted to know. So basically, if I run into any nucleotide that is not specifically specified (i.e. N, M, etc), it's automatically a mismatch. It doesn't matter if I'm comparing N with M, or N with A, etc etc etc, it's a mismatch, correct? That's all I need to know as I'm working on implementing scoring to determine similar words for alignment.


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How could I identify whether given RNA-seq data is paired end or single end
I need to have a RNA-Seq dataset and therefore, I've visited the following site NCBI-geo C. Elegans
In the Supplemantary file part, I clicked the SRP/SRP051/SRP051702  ftp and downloaded sra file. Then I need to convert it to fastq file format. For this purpose I've heard that there is a sra-toolkit and within it there is  fastq-dump exacutable. However in order to use it, first of all I have to figure out whether my sra file contains paired-end or single-end read data. So my question is that how could I know the type of reads in the dataset (in the first link) ?
Without any information, I used --split_files flag of fastq-dump exacutable and it generated two 14 Gb  (both of them  are exactly 14.346.367.840 bytes) files and their name are  SRR1741330_1.fastq and SRR1741330_2.fastq Does it mean my dataset is paired end ?
As an another question (different but related with part 1). In the SRR1741330_1.fastq file, at some lines, sequences contains different characters such as CCCFFFFFGFHHHGJJJJI#1?FEIGGI... Before looking into these files, I have thought that these sequence lines should only contain the letters of A,G,T and C . What are these F,H,J,I,#,? etc..
My questions might be trivial and unmeaningful but since I'm a totally new person in this area, I could not understand them.
EDIT
New question
I wonder one more thing, since its related with previous part of my question I asked it here instead of creating new question. After I generate fastq file with dump_fastq executable, all the sequences are length of 50. Does it because of parameters of the dump_fastq executable or its related with original sra file? In other words, could I increase this length?

To find out whether a dataset was paired-end or single-end, go to SRA, click on a run, and look under "Library". Paired-end datasets will typically have "Layout: paired". Note that people don't always mark this correctly, which causes no end of headaches.
Regarding lines like "CCCFFFFFGFHHHGJJJJI#1?FEIGGI", that's the quality score line. Look at the fastq article in wikipedia for more details.
Edit: Regarding the updated portion of your question, no you can not increase this length. The sequences produced by Illumina machines are a fixed length and that length (or whatever length was submitted) is what you're getting. As a general rule, you'll get whatever sequence was updated (unless you forget the --split3 (or whatever it's called) option).

@Devon_Ryan thanks for your help, by mean go to SRA, I think you refer to sra file that I downloaded from the link I provide above,right?

Okey, now I found it, it is in the browser

Exactly, I mean the webpage for a run on the SRA website. The same information is typically available in GEO too (click on one of the samples and read the "Data Processing" section), but it's typically a bit faster to just go to SRA.

@Devon_Ryan thanks again, could you look at my question again, I've updated  it

I've updated my reply. The short answer is that you're getting what was uploaded, which is typically all that was sequenced.

@Devon_Ryan thanks for the last time : ) So, as I understood from your updated answer, rna-seq data was supplied by the length of 50 base pairs read then.

@stackunderflow Yes, exactly.


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NCBI-geo differences between platforms,series and samples
This is the first time that I use NCBI-GEO and I've faced with 3 different categories inside it. They are platform, sample and series.
NCBI-GEO have GEO-Overview section to explain these terms to ones who are new to this website, however; although I read it a couple of times, I could not understand it totally.GEO-Overview page The terms are a bit complex for the ones who are not familiar with the concepts. Therefore, is there anyone who could explain them with simpler words ?
In my situation, I use NCBI-GEO to get RNA-seq data for the C. Elegans organism. I've downloaded the following SRA file : NCBI-geo C. Elegans sample
As far as I understood, a sample contains an individual organism's information.(In my case its a single C.elegans organism)
Series are somehow the collection of multiple samples I guess. I wonder what is the criteria to put different samples into same series?
And lastly, what is the differences between platforms and series ? I thought that each organism have a single platform, in other words, I thought I could find all the things about lets say C.elegans in its platform but I saw that a single organism type may have a lot of different platforms.
So, I'm seeking to learn these concepts and I think this website is the correct place to ask it. If my question is off topic, sorry about that I was not aware of it.

You should have a look at the overview of GEO more carefully or go through NCBI handbook on GEO

Why is this being closed for homework?

@MarchHo that link answers this completely. I VTDed because reading the help would have been sufficient for the basics. I am not too sure if VTC was appropriate in this case or not. And it is not closed yet; it requires 3 more votes.

The platform at NCBI-GEO means the particular microarray that was used (because originally most of GEO was just microarrays), OR, more recently, it could mean the type of NGS machine that was used.  For example, every microarray manufacturer creates an ADF (array data file) that maps the DNA sequence at every position on the Chip to an accession ID. And this ADF mapping file gets registered at GEO as a GPLxxxxxx. This was presumably to facilitate direct comparisons between different experiments from various labs using identical Chips (e.g. Affymetrix, or Nimblegen).
With the advent of NGS the Platform has to be a name from an approved list of companies and devices: ILLUMINA, HiSeq, etc.
A series is constructed by the data providers and allows them to cluster the results from related samples. So, for example, in the modENCODE consortium, I believe all of the C. elegans RNA-Seq from Bob Waterston's lab/Project are clustered together under one overarching GSExxxxxxx. More details at www.modencode.org and links therein.

so, every GPLxxxxx number corresponds to either a single microarray or type of NGS right ? I mean, lets assume I have a microarray. It has a GPLxxxx number and if I searched it on NCBI-GEO, I can find the experiments in which  same microarray chip was used? "Same microarray" means produced by same factory and have identical features.

@stackunderflow Check the link I mentioned. Everything about GEO is nicely explained there.

@WYSIWYG  thanks for your helps, I think I got the main idea behind these terms. Since I'm completely new in this area, some keywords are really hard for me to understand  at the first time, therefore I really appreciate your helps.


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Naming convention of miRNAs
I'm trying to understand naming convention of miRNAs. I've found the wikipedia article about it nomenclature Based on it, I try to figure out what is hsa-let-7a. As far as I understood, hsa refers to human but I expected a "mir" or "miR" after hsa but there is a word "let" .So my question is that hsa-let-7a is a name for a single mature miRNA or is it a name of a miRNA family(if there exists such a thing) ?

let-7 is a miRNA family (let stands for lethal). It was one of the first to be discovered, in fact (in C elegans). let-7a is a single member of the family. It seems rare in science that established names are changed to match newer nomenclature, though I don't know if that's the case here.

Another such example: There is an miRNA in Drosophila called bantam (Just bantam)

let-7 is a miRNA family (let stands for lethal) and, in fact, was one of the first to be discovered (in Caenorhabditis elegans). let-7a is but one of many members of the family, with orthologs in many species. It seems rare in biology that established names are changed to match newer nomenclature, though I can't say for sure if that is why many (but not all) of the let-7 family members don't follow the standard.
Edit: Coincidentally, I was just reading a review about miRNA and came cross this informative excerpt which should at least give some sense of legitimacy to my above speculation.
Ha M, Kim VN. 2014. Regulation of microRNA biogenesis. Nat Rev Mol Cell Bio 15(8):509-524

The nomenclature of miRNA genes is somewhat inconsistent. The genes found in early genetic studies were named after their phenotypes (for example, lin-4, let-7 and lsy-6), whereas most miRNAs found from cloning or sequencing received numerical names (for example, the lin-4 homologues in other species are called mir-125).

Roush S, Slack FJ. 2008. The let-7 family of microRNAs. Trends Cell Biol 18(10):505-516

The lethal-7 (let-7) gene was initially discovered as an essential developmental gene in C. elegans and, later, as one of the first miRNAs (Figure 1). Soon afterwards, a simple BLAST (basic local alignment search tool) search revealed exact matches to the mature let-7 miRNA sequence in the emerging Drosophila melanogaster and human genomes. It is now known that mature let-7 is highly conserved across animal species.
The let-7 family of miRNAs is often present in multiple copies in a genome (Table 1). To distinguish between multiple isoforms, a letter is placed after let-7 to indicate a let-7 with a slightly different sequence, and a number at the end denotes that the same sequence is present in multiple genomic locations. For example, humans have ten mature let-7-family sequences (Figure 3) that are produced from 13 precursor sequences (Figure 4). Three separate precursors produce the mature let-7a sequence (let-7a-1, let-7a-2 and let-7a-3) and precursors from two different genomic locations produce the let-7f (let-7f-1 and let-7f-2) sequence. However, the size of the family can differ between organisms. Drosophila (fly) has only one let-7, whereas the zebrafish Danio rerio has 11 mature sequences in 19 loci throughout its genome…

Early characterization of let-7:
Reinhart BJ, Slack FJ, Basson M, Pasquinelli AE, Bettinger JC, Rougvie AE, Horvitz HR, Ruvkun G. 2000. The 21-nucleotide let-7 RNA regulates developmental timing in Caenorhabditis elegans. Nature 403:901-906

Some general aspects about miRNA nomenclature in addition to canadianer's answer:

mir refers to the pre-miRNA whereas miR refers to the mature form
mir-x-y mir-x-z for example mir-1-1 and mir-1-2 refer to different precursors that give rise to same mature miRNA (i.e. miR-1). These different precursors can be regulated differently.
The mature miRNA resulting from the 5' arm of the precursor hairpin has a suffix of -5p; similarly the product from the 3' arm has the suffix of -3p. Previously, the lesser expressed/abundant (or unexpressed) of the two products was designated with an asterisk. For e.g. miR-124*. This convention is obsolete now. Any miRNA by default has the -5p/3p suffix. This is useful especially in case of let-7 because there is no uppercase notation to differentiate precursor from the mature miRNA. For example, the mature form is always denoted as hsa-let-7b-5p even though the -3p is never expressed.
A "family" consists of miRNAs with the same seed sequence (usually 2nd-9th residue). Sometimes it can be observed that one of the arms is less conserved than the other between different precursors. For example in the case of let-7 the -3p(s) do not have a conserved seed sequence, however, all -5p(s) have the same seed sequence.

And as mentioned in the other answer, the name let-7 just sticks around because of historical reasons.


text:
Is the PCT of the nephrons of the kidneys of marine vertebrates to minimize the natural tendency for exosmosis?
For marine vertebrates, the concentration of salts outside their body (in the water, that is) is more than that inside their body. So, there is a natural tendency for exosmosis to occur.
To minimise this natural tendency for exosmosis to occur, these vertebrates would need to retain more salts inside their body.
I understand that the Proximal Convoluted Tubule (PCT) of the nephrons of the kidneys is involved in selective reabsorption of salts. So, making the PCT more coiled would increase the surface area of absorption, thus increasing the absorption, and therefore retention of salts in the body.
So, my question is: is the PCT of the nephrons of the kidneys of marine vertebrates more coiled than that of terrestrial vertebrates?

What kind of marine animals are you interested in? Have a look at this post.

I've modified the question to be more specific.

Plus.. During osmosis, salts are not lost.. It is the water that is lost. The marine animals have to conserve water, not salts. You can see that other link. It is quite related to your question but is not about the kidney organization.

First off, I agree with @WYSIWYG - marine life has adapted to conserve water, not electrolytes. Osmosis in sea water draws water from the body and hence their problem is to keep the water in, not the salts.
Osmoregulation in marine mammals is accomplished mainly via differences in hormonal regulation of the kidneys. The histology and morphology of the kidneys are different, but the adaptations have more to do with adaptations to diving/pressure than anything else. Instead, most marine mammals take care not to drink too much sea water and obtain their water from metabolism and foods (Ortiz, 2001).
Reference
- Ortiz, J Exp Biol (2001); 204: 1831-44


text:
How do non-human apes cool their body temperature in warm climates?
Humans have sweat and we all know how that provides cooling along with the remaining vellus hairs acting as wicks.
So how do hominids, our close relatives (some of which are more massive than humans) regular their body temperature, given that they are completely covered in fur like typical mammals?

I’m specifically interested in how a human who had an atavism※ that’s like generalized hypertrichosis terminalis would fare and get by.
※ in my story, it’s due to a repaired chromosome 17 that didn’t turn out the way that was intended.

First they have less cooling needs than us, shady forests are cooler than sunlit savannah, they also are not as active, especially the larger ones, at least not in the hottest parts of the day. keep in mind chimps are much smaller than us so they have an easier time cooling off. They cool themselves the same way we do, by sweating, its just not s effective as it is in us, but then it doesn't need to be. .

Gorillas and urangs are larger than us.

hence needing to mention the chimps are smaller.

Some of the tactics include:
_ More frequent breaths.
_ Sweat more.
_ Loss of skin moisture.
You can read here for more details: https://physoc.onlinelibrary.wiley.com/doi/pdf/10.1113/jphysiol.1976.sp011251

Answers are expected to stand for themselves with references for further reading. It would be great if you could edit your answer with information from the paper you linked.


text:
How much penicillinase is needed to deactivate 125mg of amoxicillin?
How much penicillinase is needed to deactivate 125mg of amoxicillin?
Penicillinase in penase concentrate from CPC.
20,000 Levy Units/ml/min
I need IU of 125 mg of amoxicillin.
Please note that the problem is with finding the right parameters for amoxicillin, not penicillin G.
My goal is to test an amoxicillin product (i.e. it contains more than just amoxicillin) so the overall problem is more complex, I wanted to keep the question focused.
I would like to know the minimal amount of penase needed to deactivate 125mg of amoxicillin (that's the amount of amoxicillin in the product) in a reasonable amount of time, say 5 minutes. I want the minimal amount of penase because I am not sure what is its influence on bacterial growth, I am going to test that separately. Maybe I am going wrong about the whole problem?
I think that this paper could be helpful in general: Microbiological Applications of the Inactivation of Antibiotics and Other Antimicrobial Agents but it's behind a paywall and I can't access it at the moment. So far, I failed to find any other useful resources.

This question cannot be answered without a time factor. Assuming no enzyme degradation, theoretically a single enzyme molecule can do it given enough time.

Well, you could be a chap and answer the question, instead of downvoting, including factoring in the time factor - it would be just one abstraction higher than a simple number and also more valuable and informative.

This is not a homework question! It even doesn't fit the homework policy. The first two points are off the table immediately. The last point, "A question that addresses a basic biology concept that may seem trivial to biology professionals" is obviously invalid too since it seems none of you are able to answer the question for amoxicillin as opposed to penicillin G.

@jkbkot You should also show some basic effort. Levy Unit is not a standard enzyme activity metric. As it seems it is used only in case of Penase. Add some of your calculation on how much penicillinase is would deactivate an equal amount of penicillin. From your question it seems you are asking others to do your calculation. The edit was useful but I suggest that you add some more details.

@WYSIWYG I agree I should have added more details from the beginning, sorry about that. I'll add more of my own research a bit later in the day, thanks.

@jkbkot Moreover, there are several types of penicillinases which would ahve different activities. I don't think CPC has mentioned the composition of their product. I am afraid you would have to do an assay to determine the activity. There are no research papers (from my quick search) also that compare activities of a given penicillinase against different penicillins. Plus, the time factor is necessary. Amoxicillin can eventually degrade spontaneously as well.

@WYSIWYG thanks, that also helps - I spent quite a bit of time going through research papers myself and haven't found any useful references. Maybe one behind a paywall, unfortunately inaccessible to me at the moment. I'll update the answer

I still think this question shows an appalling lack of effort. If you had truly replicated TomD's results as above, you should have posted them in the question to begin with, so they know you have already done it and not spent their time doing your calculations for you. Also see this Meta answer for my opinion on this topic. Many members of the community vote to close questions for being homework if they are poorly researched and show no effort.

@MarchHo yes, I should have included a bit more information which I already admitted and said sorry. Also, sorry but appalling is too strong of a word and just adds more evidence to the hostility of this community. The question as it stands now is fine and it is also thanks to the effort of the helpful comments (which yours were not). If we are being pedants then everyone should have paid a lot more attention to the fact the question was about amoxicillin instead of assuming the poster is an idiot.

As March Ho has pointed out, we could do with a bit more information on this one.
I am going to attempt the calculation for penicillin G, and the necessary information seems to be available.

From here, A Levy Unit (LU) is the amount of penicillinase that inactivates 593 IU (International Units) of Sodium Penicillin G  per minute at 25°C and pH 7.0
From here and here, the specific activity of penicillin G is about 1667 Units/mg
I am assuming your concentrate is 20,000 Levy Units/ml (not 20,000 Levy Units/ml/min, which does not make sense to me)
Therefore 1 Levy Unit will inactivate (593/1667)mg, or 0.356 mg, of penicillin G per minute, pH 7.0 and 25oC
2.81 Levy Units will inactivate 1 mg penicillin G under identical conditions
351 Units will theoretically inactivate 125 mg penicillin G in 1 min under the conditions stated.
Your concentrate is 20000 Units/ml, or 20 Units per microliter.  So you will need (351/20) microlitres, or about 17.5 ul.
This assumes that the enzyme continues at a constant rate given by the specific activity right until the end.  This might not necessarily be the case, as one would expect the rate to slow down as the substrate concentration falls below Km.  Personally, I would use at least a 10-fold excess. 176ul - Seems a lot?

Please feel free to provide further information, or to edit the answer in any way you feel is necessary

Thanks for the answer! It helps me to realize that 1) I seem to understand how to compute this myself, I actually did it for penicillin G and got the same result as you 2) the real problem is that there seems to be no IU definition for amoxicillin, or at least I'm failing to find it so I'm not sure how much penase is needed. I am actually figuring out a way to test amoxicillin - maybe I could ask about that directly too? I need to double check pharmacopoeia first though...

Now I see they say that for other penicillins one should use 20 times more penase. Is there no more specific information available?


text:
genes map to eQTL/pQTL

Of the 192 target genes, 79 map to a significant eQTL or pQTL in at
  least one dietary condition.

is the line for a paper "Multilayered Genetic and Omics Dissection of Mitochondrial Activity in a Mouse Reference Population"
As far as I know an eQTL is a SNP (particular genotype) affects gene expression. How can I map then a gene to an eQTL? A gene is a sequence on a reference genome. Genotypes of one specific gene are just variations of the sequence of this specific gene. So, a gene itself cannot be an eQTL, a specific genotype (with a variation) can be, right?
So, how can it be possible that one gene (let say GENE22, location: 10-100) is mapped to eQTL(SNP within GENE22, location: 12) in one dietary condition but is not mapped to eQTL in another dietary condition if we are still speaking about the same gene and the same location of an eQTL? So, I am not quite sure what they mean or what they mapped there.

The QTL is a locus in which the genes/markers have been correlated with some phenotype. It may include markers such as SNPs but they are not SNPs in themselves. A gene can be part of a QTL, but the QTL itself is made up of  a number of markers!

"but they are not SNPs in themselves." There are studies that have lists of eQTLs as SNPs. "A gene can be part of a QTL but the QTL itself is made up of a number of makers". So, you want to say, that they had a list of eQTLs and pQTLs and they just looked in which genes they are located, right? It does not make sense for me because they are referring to dietary conditions and if the looked only at the location then there should not be differences in the conditions because location is always the same.

You take a list of SNPs and a list of gene expression values and you plug it into something like this: http://www.bios.unc.edu/research/genomic_software/Matrix_eQTL/

So, how can it be possible that one gene (let say GENE22, location: 10-100) is mapped to eQTL(SNP within GENE22, location: 12) in one dietary condition but is not mapped to eQTL in another dietary condition if we are still speaking about the same gene and the same location of an eQTL? So, I am not quite sure what they mean or what they mapped there.

The SNP is associated with expression level changes in that gene for one dietary condition but that same SNP is not associated with expression level changes in another dietary condition. Its possible that the SNP never appears in samples from the other dietary condition.

I edited my question.

so, the right way would be to say: "markers within 79 genes map to a significant ...." , right?

I'm not sure the phrasing for which is significant, my guess would be something like: "we find X significant eQTLs when testing Y genes and Z snps"


text:
Understanding recombination scoring in family pedigrees
I am having some problems understanding recombination, and I am not sure what element I am missing here. This figure is an example from my text book. The pedigree belongs to a family with an autosomal dominant disease, typed for a marker with alleles A1-A6.

So, all meioses are phase-known, meaning we know which combination of alleles was inherited from each parent. Apparently this leads us to be able to unambiguously score III1-III5 as non-recombinant (N) and III6 as recombinant (R).
This is were I get lost, I don't understand how it is possible to unambiguously score each individual as N or R. Also, if the disease is dominant, how is it possible that III6 is affected, as both alleles originate from the both unaffected I1 and II2?
EDIT: Blue colour in the pedigree means the person is affected by disease, white symbolizes unaffected individuals. I believe the red stars symbolizes that the person carries the disease gene. That would mean blue corresponds to not carrying the disease gene. If this is the case The red colour of III2 is a fault in the figure.

Could you add some details to your question to annotate the image? Also in my opinion this is quite homework question, which is off topic here at Biology SE, unless you share your efforts / research to solve the question.

I don't understand how recombination scoring works, and I thought I'd use this image from my text book to provide a basis for illustrating it. It wasn't meant as help with home work, but I understand your remark :-). How can you deduce from given alleles which is recombinant or not?

Okay so not homework :) but it would still help to know what all the stars etc mean on the image. For me at least...

To be honest I have no clue... I believe its purpose might be in emphasizing heterozygosity, but I don't think this is correct: if blue star corresponds to allel A2, then III2 and III5 should also have a blue star...

You need to add what the stars and colors are referring to.

Ns are not non-recombinant at the genomic level. What you know is that they do not show recombination events at the risk allele locus as subjects carrying the marker A1 show signs of the disease and similarly healthy subject do not carry the marker A1. Hence they are called non-recombinant. On top of that 3rd generation subjects (excepted III6) show combinations of alleles reflecting inheritance of one allele from each parent.
What is very important is that A1 is not the allele, it is the marker for the risk allele. This means that A1 is not directly the allele but a region of DNA close to the allele provoking the disease, sufficiently close to be co-inherited most of the time with the disease allele.
For III6 you actually know that a recombination must have happened as this subject obviously carry the risk allele (shows the disease) yet not the marker associated with it, therefore a recombination event happened between the marker and the disease allele.

-I've re-read the question a few more times, and what you say makes sense. I've focused too much on the picture than the text itself. My bad, guess I'm a bit tired. I've deleted my answer as yours is obviously better.

One slight remark though - On molecular level basically everybody is recombinant, but multiple recombinations can restore the original gene / allele combination, practically only those offsprings / individuals are considered recombinant who exhibit different combinations than his/her parents.

@poka.nandor You are exactly right. In the context of genetic linkage, the term recombinant is usually used at a given locus and not at the genome-wide level this is why III1-5 are considered non-recombinant, they actually are non-recombinant at the disease risk locus. Good point.


text:
Are viruses technically organisms, or not?
Are viruses technically organisms or not?
A quick Google search query for the term; are viruses organisms?, reveals various conflicting and somewhat inconclusive arguments & opinions from a range of sources. (Examples provided below).

Note: These answers are mostly opinion-based and inconclusive; falling short of the Stack Exchange community's quality control standards.

Let me be clear: I am not seeking opinions, or debate; but rather definitive answers supported by facts.

Wikipedia | Organism.
  The most common argument in support of viruses as living organisms is their ability to undergo evolution and replicate through self-assembly...
  Some scientists argue that viruses neither evolve, nor self- reproduce.
~ https://en.m.wikipedia.org/wiki/Organism
Encyclopedia of Life | What is a Virus?
  A virus is a microscopic organism that can replicate only inside the cells of a host organism...
Viruses infect all types of organisms, including animals and plants, as well as bacteria and archaea.
~ http://eol.org/info/458
Virology Blog | Are Viruses Living?
  Therefore, viruses are not living things...
  So technically speaking they are non- living organisms.
~ http://www.virology.ws/2004/06/09/are-viruses-living/
Popular Science | Are Viruses Alive? New Evidence Says Yes.
Viruses seem to carry out only one life process, reproduction, but even then, individual viruses don't carry translational machinery, namely, the proteins needed to read their DNA and RNA and build new viruses. They invade a cell and hijack its genetic tools to do it for them...
  But within the last decade, developments in virology have started to reveal more and more that viruses might in fact be alive...
One was the discovery of mimiviruses; giant viruses with large genomic libraries that are even bigger than some bacteria. (To put this in perspective, some viruses, like the Ebola virus, have as few as seven genes.) Some of these giants have genes for the proteins that are required for translation—those readers of DNA and RNA that in turn build new viruses...
  This throws the lack of translational machinery argument for classifying them as nonliving on its head.
~ http://www.popsci.com.au/science/medicine/are-viruses-alive-new-evidence-says-yes,409690
Science Daily | Virus.
  A virus is a microscopic particle that can infect the cells of a biological organism...
  At the most basic level, viruses consist of genetic material contained within a protective protein coat called a capsid; the existence of both genetic material and protein distinguishes them from other virus-like particles such as prions and viroids...
  It has been argued extensively whether viruses are living organisms...
Most virologists consider them non-living, as they do not meet all the criteria of the generally accepted definition of life...
  They are similar to obligate intracellular parasites as they lack the means for self-reproduction outside a host cell, but unlike parasites, viruses are generally not considered to be true living organisms...
  A primary reason is that viruses do not possess a cell membrane or metabolise on their own - characteristics of all living organisms.
~ https://www.sciencedaily.com/terms/virus.htm

@Remi.b Thankyou. It is worth linking, however; not the same question. Please don't report it as a duplicate. Cheers.

@tjt263 Can you explain how this question is different?

it is the same question, hence you've linked to sites that ask are viruses alive?

If you consider the definition of an organism to be 'something alive', they're the same question; if not, they're not.

So, if you think about where organism actually came from, organization, the virus is actually quite a very organized little compartment of stuff (without going into details). There are elements that have to cooperate within the actual virus to get anything done, thus, I would qualify that yes viruses are organisms. You wouldn't consider something like a prion or a viroid an organism, however, because on their own there's really just one part with no interdependencies.

Definitive suggests by definition. By definition, no. Fact- a virus is not an organism by definition. If the definition changes to include viruses, then yes. But for functional purposes, we speak english- which means utilizing the definition of words. The reason why a virus is not an organism is because it requires a host organism for replication. DNA is also not an organism.


text:
Taxonomy: Categorising Collembola
Observe: In some rows (Order, Family), the third column reveals a discerning physical characteristic or body type (Elongate, Slender).
What are the discerning characteristics, missing from the other rows (Superfamily, Subfamily, Genus, Species)?
Basically I want this knowledge so I can work towards being able to separate, categorize (catalogue) and/or identify new or unknown species, etc.

Imagery
Exhibit A

Entomobrya unostrigata
(aka. Cotton Springtail)

Taxonomy
Domain      | Eukaryota        | (Eukaryotes)
Kingdom     | Animalia         | (Animals)
Phylum      | Arthropoda       | (Arthropods)
Subphylum   | Hexapoda         | (Hexapods)
Class       | Collembola       | (Springtails)
Order       | Entomobryomorpha | (Elongate)
Superfamily | Entomobryoidea   |
Family      | Entomobryidae    | (Slender)
Subfamily   | Entomobryinae    |
Genus       | Entomobrya       |
Species     | Unostrigata      |

Sources:
Image; photographed by myself. Text; transcribed from my own hand-written notes. Most likely (originally) gleaned and adapted from publicly available, online information. It may contain errors or inaccuracies.

Resources: Check out Bugguide.net and Collembola.org.

Reprise
Okay, the question has been put on hold, preventing answers from being posted (very irritating and obstructionist policy for those seeking answers, I might add). So I will attempt to clarify.

Exhibit B

Orchesella cincta

Exhibit C

Willowsia nigromaculata

Exhibit D

Entomobrya multifasciata

Exhibit E

Seira bipunctata

The diversity of these specimens; span not only multiple species, but multiple genera. Yet, despite the fact that they each exhibit relatively distinctive combinations of patterns and colours across their cuticle (epidermis, surface epithelium), setae, scales, etc. they all look pretty much identical.
This seems to be the equivalent of declaring a homo sapiens of African descent; a different species to a homo sapiens of Irish descent, on account of their superficial aesthetic differences (most notably regarding melanin; the pigments that determine the colours of a human's skin, hair[s], and iride[s] (eye[s]), and so on). Which, at first seems absurd. But for those of you who disagree, observe the differences between (for instance): Exhibit X vs Exhibit B & Exhibit Y vs Exhibit E. Similar variations between individuals of the same species are not uncommon in the least.
Thus, my question: What are the discerning taxonomical differences between various species and genera of the Arthropoda known as Collembola (aka Springtails)?

Exhibit X

Orchesella cincta

Exhibit Y

Seira bipunctata

Don't know the answer to your question, but I have had the same problem as you; feeling like there should be a set of criteria for each and every binomial systematic group (otherwise what's separating them?). Also, I sometimes find that once you get to Order, Superfamily, Subfamily, Subfamily, they groups tend to change from resource to resource making them unreliable, there doesn't seem to be any one official resource of all the known species in all of their groups. So, yes, I agree, being a bit of a perfectionist myself I find it annoying that there isn't always a criterion for every group.

Is the third column supposed to contain characteristic features? At the moment, this is not really the case (its a mixture of common names of taxonomic groups and physical descriptions). I also doubt that "slender" can be used by itself as a trait to separate Entomobryidae from other families. Are you looking to construct a dichotomous Identification key of springtails, or what is the exact purpose of the question?

Continued; If the purpose is a determination key, you need to decide whether your key is supposed to mimic the phylogenetic situation (which your table implies), or if it supposed to be easy to arrive at a species determination. For a key that moves along with the phylogenetic tree, you will need to find good "taxonomic characters" for all taxonomic levels, that are easy to diagnose. If you are only looking for a way to determine species, there might be better ways to structure the key (even though it is in many situations preferable that keys follow the phylogeny).

The third column is just for notes, including common names, etc. But if you're really getting hung up on that, you're probably missing the point of the question.

@LucasDyson-Diaz is on the right track. Some kind of key or matrix would be nice, for instance; a discerning characteristic of arachnidae VS insecta, might be something like 8 legs VS 6 legs, or something of that nature.

Yeah @tjt263 , also I've tried to make a chart, as you describe, but I've only succeeded to make it to phylum level since there are many in the prokaryotic kingdoms, still to that level it's difficult to distinguish organism from organism at such a high (phylum) level let alone order, class, etc.

The third column doesn't contain distinguishing characteristics at all -- they are the first word of the vernacular (or common) names for those groups on Bug Guide. Specifically, Bug Guide lists order Entomobryomorpha as Elongate-bodied Springtails, and family Entomobryidae as Slender Springtails -- so I think you're just looking at truncated data that's misleading you!
What you're looking for a taxonomic or identification key, which gives you set of yes-or-no questions that will allow you to identify an individual to a particular taxonomic group. Know Your Insect has a taxonomic key that allows you to identify insects to order Collembola, which provides exactly the sort of distinguishing characteristics you're looking for: "This illustration shows two characteristic features of members of this order: the collophore (center), a tube-like structure used to secrete fluids; and the furculum (lower left arrow)." Note that this resource refers to this group as an order, not a class -- that's not uncommon among different taxonomic resources. Wikipedia refers to this group as subclass Collembola!
Unfortunately, it doesn't look like there's a freely available taxonomic key for species within this group, unless you buy a specialized guide like A Key to the Collembola (Springtails) of Britain and Ireland.

You're wrong; elongate and slender are discerning physical characteristics. For example: The Poduromorpha and Entomobryomorpha have an elongated body, while the Symphypleona and Neelipleona have a more rounded, globular shaped body. But yes, the taxonomic rank of Collembola has been disputed, given the more recent genetic evidence that suggests any similarities between springtails and insects are a matter of convergent evolution.

Sure, but they aren't very helpful characters (how slender? Elongate how?), and the first couple of values are clearly not characters at all (What sort of characters are Arthropods? Springtails?), which is why I'm pretty certain that third column is common names, not characters. I think you're looking for are apomorphies -- characters shared by individuals within a group but not found within its close relatives, like hair in mammals or hollow bones in birds.