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How can one measure the effect of a stem cell's matrix on the progeny of that stem cell?
A number of recent papers that have used hydrogels to alter the ECM of mesenchymal stem cells to quicken their development to cells used specifically for cartilage or really any other type of cell.
Here is an example of one of the articles
However, even after looking at the data sets from all of these experiments and research projects I still don't understand how this is done (i.e. what lab instruments, the actual procedure), for example of cartilage progenitors from stem cells.
I specifically want to know how this occurs in neuronal stem cells and how can we isolate those stem cells and test the effect of encapsulating them with a type of hydrogel to alter their fate? How could this be done?

I made some edits to your question for clarity - please check them out and alter if necessary. Thanks for your contribution!

The stem cells respond to the force they exert to the surface that they are plated on. Very soft matrices mimic the environment of the CNS resulting in neurons and hard matrices will result in cells like osteoblasts as this is more representative on the microenvironment of bone. The tensile force changes the expression patterns of the stem cells causing them to "choose" a particular fate. As for testing what is what, the way that I have seen is to do RNAseq on a sample of the cells and look at their expression patterns. Neural progenitors would be identified by expressing certain genes

Thank you, both of you. @AMR After looking into soft and hard matrices and how they actually affect a stem cell's fate, I found that it was indeed one of the ways you may motivate a stem cell to turn into a certain cell. I believe in my research I will be specifically searching for a way to chemically alter the surroundings of the stem cell to change it. But the advice about the RNAseq is great, and I will definitely be looking into that.


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What are chemical candidates for replicator molecule?
I have read Selfish gene by Richard Dawkins and idea is that at random some molecule was synthesized that had a property if there is enough 'materials' to construct copy of itself - it would. And then it made bunch of it's copies and then copies started mutating and eventually we got to DNA in us.
In his book he never says what actual molecule is original replicator.
What are chemical candidates for original replicator molecule?

As @AMR said, there are self replicating ribozymes. That's something that we know. There can be several such candidates, theoretically.

There is no accurate answer  , as you say we talk about candidates . there is lots of strong theories about the origin of self replicators . RNA world was a strong candidate since latest experiments that suggest there is a good chance that some molecules that have simpler structure than RNA , could have a chance to do self replications .
    there is also strong theories such as :
1.Prions that delivered by space objectives and Meteorites .
2.Life's origins may result from low-energy electron reactions in space:
http://phys.org/news/2016-06-life-result-low-energy-electron-reactions.html

first self-replicator recreated in lab:
RNA, or something very like it, has long been a strong candidate as the first self-replicating molecule in the origin of life. This is because it can both catalyse chemical reactions and carry genetic information.
But chemists first needed to explain how a large, complex molecule like RNA could form spontaneously to begin the process. They had done so for some, but not all, components of the RNA molecule.
https://www.newscientist.com/article/2088006-building-blocks-of-lifes-first-self-replicator-recreated-in-lab/
The origin of replicators and reproducers:
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC1664675/
you can also check the topic :The RNA World and the Origins of Life
from the book :
Molecular biology of the cell by Alberts B, Johnson A, Lewis J, et al.

after all this article was so innovative for me . I think it's a good explanation for self replication of molecules simpler than RNA.
http://www.nanowerk.com/nanotechnology-news/newsid=40896.php
"Maslov and Tkachenko's model imagines some kind of regular cycle in which conditions change in a predictable fashion—say, the transition between night and day. Imagine a world in which complex polymers break apart during the day, then repair themselves at night. The presence of a template strand means that the polymer reassembles itself precisely as it was the night before. That self-replication process means the polymer can transmit information about itself from one generation to the next. That ability to pass information along is a fundamental property of life"

The details appeared in a paper titled “Prebiotic Phosphorylation and Concomitant Oligomerization of Deoxynucleosides to form DNA.” The paper’s authors say that their new finding is the latest in a series of recent discoveries pointing to the possibility that DNA and its close chemical cousin RNA arose together as products of similar chemical reactions, and that the first self-replicating molecules—the first life forms on Earth—were mixes of the two.

https://www.genengnews.com/news/rna-dna-world-circumvents-rna-world-sticking-point/

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In Darwinian evolutionary theory, does the environment has any role?
In Darwinian evolutionary theory, does the environment has any role?
To state it clearly:
Suppose I have an isolated system. First, I place several types of entities (organisms) in an environment. According to Darwin, eventually, the fittest will survive.
Next time, I place Only one organism. Will it survive this time as there's no one else in the system?
If not, did Darwin also consider the whole environment as another entity, so that in the above case, there are two entities, the organism and the environment. The environment is more evolved and it survives, other dies?
Moreover, if the climate is favourable and the organism reproduces sexually results in the same fate as the case if climate is unfavourable, organism reproduces asexually, what conclusion does it leads to?

For a great introduction to modern evolutionary theory, check out Understanding Evolution from the University of California, Berkeley. Darwin (and Wallace) may have originated the first coherent theories of evolution, but things have changed a lot since then. The environment absolutely has a role in evolution, otherwise there would be no stimulus to evolve. Additionally, there's no such thing as a truly isolated system - predators need prey, all depend on plants and/or other organisms that depend on the sun, etc. etc.

I don't think you understand Darwin's argument very well.  Organisms never evolve.   If you place one organism, there's nothing to evolve, so your question is meaningless.  Populations evolve. If the organism produces a population (i.e. through its progeny) then there is room for evolution on the population.

Those best adapted to their environment will reproduce. Fitness is a value judgement. Take a human with sickle cell anemia and place them in the Himalayas and they will suffer terribly and likely die before they can reproduce. Place the same person in a jungle type environment, and maybe they are better adapted to survive until they reproduce as they are resistant to malaria, where as a Sherpa might quickly fall ill in the jungle and die from trypanosomiasis before reproducing. Both organisms are "fit" just in different ways that the environment places pressures on.

Fitness has no meaning without the environment. There is no absolute fitness. Fitness is always with respect to a given environment.

"According to Darwin, eventually, the fittest will survive"
Darwin never said this. Fitness and survival are two separate concepts.
In a simplistic version, fitness is measured by reproductive output. Survival is, well, survival. You can have higher output than your brother, but have a lower chance survival. In fact, the very act of producing offspring can negatively affect your ability to survive.
No, the environment is not an other organism, in any context. It is the place (stage, arena, etc) in which the measure of fitness (reproductive output) can be measured. The environment can directly influence and affect fitness, and other organisms (inter and intraspecific) ARE part of that environment.
A single isolated organism does not evolve, but can have offspring. However, you need a more than one organism for fitness to have any meaning in a Darwinian model.

It would have to be a bacterium or "simple" eukaryote that either replicates asexually or through self fertilization in order to have offspring if they are isolated. Mammals, reptiles etc. need certain levels of organisms in a population to allow for successful enough reproduction to maintain the species above extinction.

Yes, but it makes do difference to the central concept of fitness vs survival. I did not qualify the organism in isolation.

Prokaryotes and viruses (even prions) can have fitness. Anything that replicates with information persistence can have Darwinian fitness.

-1: I don't think your answer is particularly clear or well constructed, has no references or supporting material, and generally does a poor job of answering the question.

Fitness is a function of both reproductive success and survival. The answer is wrong.

Sorry, Remi.b, but your understanding of fitness is elementary school, at best. Fitness has never meant survival. Without offspring, your fitness is zero. Period.

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Allele-specific bisulfite sequencing
When evaluating methylation status at various CpG sites after sequencing, how much consideration should one give to random single base pair insertions and deletions. Suppose there is a CA dinucleotide; can we assume that the CA is native to the sequence or results from a G deletion especially when the latter is suspect once comparing to other sequences. Is there really a set standard sequence to compare it?

Welcome to BiologySE! Thanks for your question...can you give a little more detail about your question, specifically the rationale behind it so other individuals can follow along and better understand what you're asking? You may want to check out the help center as well, for some great tips on how to ask a great question... http://biology.stackexchange.com/help

Are you talking about when a single read in your deep-sequencing experiment has differences in base composition compared to other reads mapping to the same locus?  Are you suspecting a duplication event where the original copy is CpGpA and the copy you are referring to is CpA?

I agree with Vance, we need a little more detail to better answer your question. From what I can tell, you are asking whether a single nucleotide polymorphism (SNP) that results in the addition of a cytosine base to your sequence is of concern when examining the methylation signature of that sequence.
I would first determine whether the polymorphism is common in the population using NCBI. The more common the polymorphism, the less likely it is to have a serious effect. However, that is not to say that it doesn't have any effect.
I would then examine how much the methylation of that one site varies compared to samples without the additional cytosine base.
Finally I would determine the importance of the location of the SNP. Is it located in a CpG island? Or is it located on the shores, shelves or 'open sea'? (see work by Sandoval et al.)
Without knowing all the above it is hard to make a call on how important/unimportant the SNP is in your methylation analysis.


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Membrane resistance of a neuron
I am about to simulate a neuron activity with the "Leaky Integrate and Fire" neuron model. But for that I need the membrane resistance. I was really looking a lot online, but I just cant find a value. Does anybody know the measured resistance value, or the value usually used in this model?

Welcome to BiologySE - thanks for your question... do you think you could give some references to the "Leaky Integrate and Fire" model to help other individuals better understand your question? Also, you may want to check out the help center for pointers on how to ask a good question...http://biology.stackexchange.com/help


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DNA Sequencing Types & Qualities
I've been searching for a list of the types of DNA sequencing (e.g. Sanger, Next-Generation) and how prone they are to sequencing errors (None, Somewhat, Very), but I haven't been able to find anything.  Here are a few scenarios:

I have a bacterial culture I want to sequence to see if a plasmid was properly transformed.
I want to verify a PCR product.
I want to sequence a large organism's genome.
I want to verify a mutation found in a human tumor (found using exome data)

Given these examples, and any other you use regularly in the lab, can you please give me information on overall platforms (not specific like Illumina, but general like Sanger sequencing) and their error prone-ness?

Don't you think your question is a bit broad? Did you check out reviews on this topic? There are many. See these: http://dx.doi.org/10.1101/gr.168393.113, http://dx.doi.org/10.1186/1471-2105-13-185, https://www.biostars.org/p/2015/

I'll be honest, this seems like an exam question reworded.

@James It's 3 years old, but definitely wasn't an exam question. I was a PhD student and was getting started in bioinformatics & my uni's resources to inform us on this tech was underwhelming. I was trying to figure out what tech I needed for different experiments based on their error rates.

In that case, I think it suffers from needing to be more focussed like @WYSIWYG suggested originally. Essentially this would be more helpful if it was split into 4 questions.

Sequencing does not seem necessary for this... most plasmids contain an antibiotic resistance gene (ex. kanamycin-resistance) that will allow cells that have been appropriately transformed to survive in the presence of kanamycin.
If it's just a PCR product, Sanger sequencing seems the best way to go. It is quite specific and not prone to algorithmic errors as are next-gen sequencers.
Next-gen sequencing would be ideal as you're dealing with the whole-genome. Sanger would prove laborious and low throughput. Any variants of interest you find call always be confirmed through sanger. Illumina sequencers are quite robust at sequence generation at the whole genome level.
This mutation can be verified through sanger sequencing, but you could also conduct targeted re-sequencing of the particular exon (although this seems like over-kill). Since it is a single mutation, I believe the specificity of sanger would be your best bet.


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Why do I drop my pen in a drowsy lecture?
I know that there is loss of muscle tone during REM sleep. Is this loss in muscle tone the reason I generally drop my pen in a sleepy lecture? But we were taught that sleep has 4 stages and REM sleep occurs after the 4 stages, ~90mins(?) post sleep. So technically, it should be that I'd be dropping my pen after 90mins of sleep. Though this is not the case. The pen drops generally when I'm at verge of sleeping.
I realise that there also is muscle atonia during other stages of sleep but, there is this famous story of Salvador Dali and the way he slept. He had a key in his hand and kept a tin plate beneath, as soon as he reached REm (dream) sleep, the key would fall and he'd wake up to draw stuff that he saw in his dream.
Why did Dali's key drop at REM sleep but my pen, way before that?

Is "dropping the pen" just a figure of speech?

@wysiwyg No! I mean, the pen falls from my hand, only to wake me up from sleep

I meant to ask if your question has something specific about the pen. What if you don't hold a pen while you go off to sleep during your lectures?


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What is the conical mouthpart of a soldier termite and what does it do?
I'm trying to get a visual understanding of termite anatomy and I'm getting quite confused on the protruding mouthpart area (it looks like a beak) located between the pincers.
What does it do / what is it for? It does not look like a chewing mouthpart.
Further, is it related to the mouthparts of the worker / females termites?
Oddly enough, I found "Conehead" termites with what appears to be a far more profound version of this part (though it might not be) and again I see no purpose for it.
Is there a place on the internet where the breakdown of termites can be found? This would be very useful.

What is between the pincers?

This is actually the termite's labrum. Its quite big in termites. Does not have value in telling male from female.

Hi Jules~verne. We're looking for long answers that provide some explanation and context. Don't just give a one-line answer; explain why your answer is right, ideally with citations. Answers that don't include explanations may be removed. Please update. Thanks! :)


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When was it discovered XY chromosomes decide the sex of a child in humans?
Several stories are told from before-genetic-age (books and movies are my reference, the one present in my mind is Marie-Antoinette by Copola) in which we can always see that women are blamed for giving birth to a girl - when it was a boy which was "required".
How did mankind's (man) society took the discovery of the fact that the sex of a human baby is decided by the Y or X chromosome carried in the masculin genes within the sperm fertilizing the female's egg. (at least this is what I learned in school decades ago).
When was this fact discovered and by whom?
(The occasion for this question is Nettie Stevens 155th birthday today)

For history of science questions there is another website: History of Science & Mathematics SE.

The discovery of genetic sex determination, and determination of sex via male gametes (in XY species, female in ZW), occurred over some time in the late 1800's and early 1900's. Advances were made with methods to stain chromosomes and, in 1891, Henking noted that wasps produced sperm with a varying number of chromosomes. However, he was unable to gather evidence that the "x-element" was determining sex. Nettie Stevens, in 1905, then used beetles and noted an odd set of chromosomes in sperm, which later became known as the sex chromosomes. This was the discovery of sex determination through male gametes.

"In 1905, while studying the gametes of the beetle Tenebrio molitor, Stevens noted an unusual-looking pair of chromosomes that separated to form sperm cells in the male beetles. Based on her comparisons of chromosome appearance in cells from male and female beetles, Stevens proposed that these accessory chromosomes were related to the inheritance of sex." - Link

In 1905, Beecher-Williams also made the same discovery, so they are both credited with discovering sex determination by the sex chromosomes. However, Clarence Erwing McClung also contributed to theory that Henkings X-elements may determine sex in 1901.
Thomas Hunt Morgan also contributed some major work on heredity and sex chromosomes, having been initially sceptical of Stevens' discovery. He found that eye mutations in Drosophila appeared to be sex linked, and concluded that the mutations were likely carried on the sex chromosomes.
There is a paper from 1910 where Michael Guyer concludes that accessory chromosomes (the sex chromosomes) likely determined sex in humans.

"It is probable that in man and certain other vertebrates, as in the insects, myriapods and arachnids, the accessory chromosomes are in some way associated with the determination of sex."

However, Theophilus Painter, in 1921, concluded for the first time that human sex determination was by the presence of X or Y in the sperm; Guyer had gone for an XO system. While Painter got the chromosome count wrong, being hampered by the techniques of the day (an issue that wasn't resolved until the 1950's), he got the sex determination system right.


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Can salt-water frogs survive in normal water (A well)?
I don't know much about frogs, but there seem to be salt water frogs.
1) Are there such things as salt water frogs?
2) Are there frogs that live in the sea? (in reference to Vivekananda's quote)
3) Can salt water frogs survive in well water or fresh water?

Well, one day another frog that lived in the sea came and fell into the well. - Vivekananda

There are no salt water frogs. There is only one species which lives in mangrove swamps and is perhaps the only amphibian that can tolerate salinity.

There's a bit more detail on the crab-eating frog here: http://www.biodiversityscience.com/2011/04/27/saltwater-frogs-water-conservation/

@WYSIWYG So, was Vivekananda's quote "A frog that lived in the sea" scientifically wrong. Also see the updated Q.

@Wally He must have meant it metaphorically.

He also mocked evolution so.. http://www.swamivivekanandaquotes.org/2014/04/swami-vivekanandas-frog-in-well-story.html

"A frog lived in a well. It had lived there for a long time. It was born there and brought up there, and yet was a little, small frog. Of course the evolutionists were not there then to tell us whether the frog lost its eyes or not," - Vivekananda

frogs are a pretty specialized/derived group in amphibians anyway so it is not surprising they are fresh water only, there are saline living salamanders though, well one species anyway, Ambystoma taylori.

According to Wikipedia the Crab-eating frog is the only amphibian that can survive in sea water. However Wikipedia also suggests that it can only survive brief excursions into the sea.
There are no such things as sea frogs.
Even the Crab-eating frog doesn't live in the sea though. Its natural habitat is mangrove swamps.

The crab-eating frog (Fejervarya cancrivora), is a frog native to south-eastern Asia including Taiwan, China, the Philippines and more rarely as far west as Orissa in India. It inhabits mangrove swamps and marshes and is the only known modern amphibian which can tolerate brief excursions into sea water.
This frog can tolerate marine environments (immersion in sea water for brief periods or brackish water for extended periods) by increasing urea production and retention, and by remaining slightly hyperosmotic within urea and sodium flux. Adults can survive in salt water with salinity as high as 2.8%, and tadpoles can survive salinities as high as 3.9%.

https://en.wikipedia.org/wiki/Crab-eating_frog


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Do male humans have a mons pubis?
Wikipedia writes:

In human anatomy, and in mammals in general, the mons pubis (also known simply as the mons, and known specifically in females as the mons Venus or mons veneris), is a rounded mass of fatty tissue found over the pubic symphysis of the pubic bones. In human females, the mons pubis forms the anterior portion of the vulva. It divides into the labia majora (literally "larger lips"), on either side of the furrow known as the pudendal cleft, that surrounds the labia minora, clitoris, urethra, vaginal opening, and other structures of the vulval vestibule.

The Foundational Model of Anatomy suggests that the feature also exist in the non-gender specific pelvis concept. Does it?

Technically men do have a mons pubis but it is less pronounced. With increased estrogen in females it becomes more pronounced.
Wikipedia

Please add some references to your answer.

pronounced = expressed?


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How well can proteins discriminate between ATP and GTP? Can ATP act as a GTP mimetic?
GTP and ATP are similar structures with the adenosine and guanosine groups differing. Both are involved in a vast array of biological functions.
However it has been shown that in certain cases, GTP and water can mimic ATP (Niefind et al., 1999).

Are there examples where ATP has acted as a mimetic of GTP? Or in other words, are there any proteins that can't discriminate between GTP and ATP very well?

There are certainly some enzymes that don't discriminate (well) between the two purines. Is that what you mean by mimic?


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How to assess if biological measurements follow a normal or a log-normal distribution
I am using a dataset composed by $m$ samples and $n$ features (genes). Each data point is real number.
I want to understand how to preprocess data before analysis, in particular: do data points follow a normal or a log-normal distribution?
I thought about using qqplots and searching for different tests to assess the form of the distribution, but I have a doubt:
Do I have to assess the form of:

each sample distribution
each feature (gene) distribution
the whole dataset ($m$ samples x $n$ features (genes))

?

This might be better suited for [stats.se]

@C_Z_ you are right, but I thought it may be a task well-known for bioinformaticians, e.g. while using microarrays. However, if not, how can I transfer it to CV? Thanks

Have a look at http://biology.stackexchange.com/q/37167/3340

From personal experience, nearly all count data whether from microarray or reads from RNAseq of some kind, requires a log transformation of the counts. Usually a small fraction is added to all values before doing so to zero protect. Log2(counts + 0.5) or some such.   This is independent of the treatments. If you log transform one sample, you will do the same for all samples. To examine for normality, a simple way is to look at the histogram of counts (by all samples or by each sample) before and after transformation. Roughly bell shaped -> proceed.
Pictures below from my data. Although the data are from RNAseq, microarray data should be similar.
R code here:
hist(t$counts,breaks=100,main="Histogram of Raw Counts from RNAseq")
hist(log(t$counts + 0.5,2),breaks=100,main="Histogram of Log2
transformed Counts from RNAseq")

There are ways to statistically check for normality, and I find visually examining the data as I listed above is a good first step.

Preprocessing will and should always depend upon the biology that you try to answer or discover (e.g.: There might be an experimental rationale to believe that some genes behave differently in individual samples - and that different samples could possibly have different distributions.)
log-transforming your data by itself is usually no problem, and hugely facilitates the simultaneous exploration of different magnitudes (though adding a small value prior log can soon make your analysis misleading, if you intended to quantitatively study variance across samples)
For testing normality you might want to apply the Lilliefors test on raw data and log-transformed data
If you are using a readout of gene-expression you must not anticipate a unimodal distribution, e.g.: Metazoans have two different classes of genes, which overall lead to a bi-modal distribution (Hebenstreit et al. 2011) (If you can fit any unimodal  distribution - such as lognormal - you should become very suspicious and check the quality of the experimental data.)


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Could we restore some animal's muscle structure by it's bone?
Suppose that elephant dies out one day, nothing left to researchers but the fossil. Could fossil infer the existence of large nose?
If there is possibility, how exactly should they perform the deduction?

I think you would be surprised to know that forensic experts use the skull and face bone structure of a dead body to predict how the person might have looked, probably by getting the muscle structure from bone structure. I don't know whether it's true, I heard it in a TV show.


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Passing on Alleles
Just a basic question on alleles, I am just trying to learn the basic concepts.
So if we are to screen two parents, both with one dominant and one recessive allele for a given characteristic e.g. eye colour, does this mean that we can say for certain (given a correct screen) that the child has a 3/4 chance of having the dominant characteristic (expressed) and a 1/4 chance of having the recess characteristic?
Similarly, if both parents had 2 dominant alleles, does this mean it is certain for the child to exhibit said quality given that the screen was effective?
Also, are there only these two types of alleles? I heard something about gene driving creating 'super dominant' alleles and would appreciate if someone could explain this to me.
Finally, is it the case that all alleles have a 50% chance of being passed on?
(assume in this question there aren't any relevant mutations going on)

It can be a little confusing. A genetic allele is just a portion of the genome responsible for an observable trait or phenotype. Examples of phenotypes consist of your example (eye color) and millions of others. As @swbarnes alluded to, most phenotypes are complex phenotypes, meaning they are not described by a single genetic locus. Height, weight, hair color, pulmonary function, and eye color are all examples of complex phenotypes (or traits); meaning that more than 1 genetic locus controls the observed phenotype. Other traits like PTC tasting bitter are determined by a single genetic locus; these are referred to as simple traits.
When it comes to dominant verses recessive, this classification is usually applied to simple traits, and refers to whether it requires two copies or one copy of a causal genetic variant to observe the phenotype. For example, a single copy of the genetic variant responsible for sickle cell disease does not cause the phenotype of sickle cell anemia. It takes two copies to cause it. Thus we label this genetic variant as a recessive allele. If a single copy of the genetic variant were sufficient to cause the observed phenotype, then we would label that variant a dominant allele.
Some traits are intermediate, in that a single copy of the allele causes an intermediate phenotype, that is more exaggerated if you inherit two copies. The PTC tasting bitter above is such a trait. If you have a single variant allele, it will taste bitter, if you have two copies of the variant allele, it will taste very bitter. If you have zero copies, you will wonder why your friend with two copies just made a horrible face.
IF you are referring to a simple trait (controlled by only one genetic region) and the allele controlling the observed trait is a dominant allele, this would mean a single copy of the variant allele is sufficient to see the trait. However, this also means that both parents could be heterozygous for the allele ( each possessing 1 copy of the variant and one copy of the normal). In this case, there is a 25% chance their offspring gets both normal alleles. So your supposition was correct if the trait is simple and dominant.
With regards to 50% chance of passing any given allele, yes. However, in complex traits, the math gets complex quick with regards to odds of any given phenotype.
Lastly, some traits are influenced by both a genetic determinant and an epigenetic determinant that can be environmentally driven which provides infinite more complexity. I'll let look you look that up or ask another question :)  The same with "gene drives".
A neat listing of simple traits in humans and whether they are dominant or recessive comes from here:

does this mean that we can say for certain (given a correct screen)
  that the child has a 3/4 chance of having the dominant characteristic
  (expressed) and a 1/4 chance of having the recess characteristic?

If you are sure that the phenotype is 100% determined by that allele only, then yes, barring things like new mutations.  Biology always has some exceptions.

Also, are there only these two types of alleles?

In real life, most characteristics are not well described with simple Mendelian genetics.  Most traits are controlled by many genes, and the different alleles can interact in many potential ways.

Finally, is it the case that all alleles have a 50% chance of being
  passed on?

In general, yes.  Meiosis puts half of your alleles into half your gametes, and half of your alleles in the other half.

Thanks
I understand that most characteristics (phenotypes?) will rely on a number of different types of genes and the exact types may not be understood as of yet. However for characteristics which are only affected by a single gene i.e. eye colour (?) does this mean that such alleles can only be recessive or dominant?

Further, am I correct in saying that there are more than two possible types of each gene and that as a result of mutations there could be very large number of alleles? For each such allele, if it is say dominant or recessive, are all dominant genes 'equally dominant'?

Eye color is affected by multiple genes, though if a particular family has only brown and blue eyes, that variation is likely caused by variation in a single allele, and should mostly behave in a Mendelian manner.

We label one allele dominant to another if the heterozygous condition looks the same as the homozygous condition for the "dominant" allele. Like blood type; the AO phenotype is identical to the AA phenotype.  A is classically dominant to O.  But if a snapdragon with red flowers is bred with one with white flowers, the result is pink flowers.  Neither allele is classically dominant to the other.  People with AB blood type have a different phenotype than people with A or B blood.

Just because 95% of your genetics homework involves traits that fit the Mendelian paradigm doesn't mean that anywhere near that much real biology does.  Most alleles and traits simply do not fit the classical Mendelian paradigm.  The results are more complicated to predict than "This allele always masks that allele"

I understand that the vast majority of traits can not be characterised under one gene.
Is there a more complex mechanic behind it, or would it say require information from n different genes (which would mean we could theoretically calculate it)?

I think the "gene drive" that you are talking about involves the CRISPR system, and it is an engineered run of code that deliberately makes that allele 'super dominant' (by making more CRISPR components that are loaded with the particular gene replacement instructions), like in a scenario where they are trying to replace the faulty gene for Huntington's disease or ALS in an adult.


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Enzyme Activity under High Pressure
Wikipedia has this image titled "Pressure Chamber to measure Enzyme Activity under High Pressure"
Made me wonder why do we need to measure Enzyme Activity under high pressure?
Are there enzymes in nature that must work under variable pressure conditions? My impression from my knowledge of conventional chemical kinetics was that in the liquid phase the effect of pressure on chemical kinetics was typically very low simply because of incompressiblity of liquids. Unless Pressures were very very high. Is this not so with enzyme activity? Or are there enzymes that must work under very high pressures?
https://en.wikipedia.org/wiki/Enzyme_assay

Not sure what this device is, but I would guess pressure is important mostly for enzymatic reactions that involve dissolved gases (O2, CO2, etc). Increasing the gas pressure above the liquid would serve to raise concentrations of dissolved gas.

At least one group who have published on the effect of pressure on enzyme activity (Cho and Northrop) are not interested in it from a practical point of view, i.e. they are not subjecting enzymic reactions to pressures that they think they encounter naturally. Instead they are using the effects of pressure to test the kinetic and thermodynamic predictions of different models of enzyme catalysis. In particular they believe that pressure effects provide a means of testing the idea, first suggested by Linus Pauling, that an enzyme has a greater affinity for the transition state, rather than for the substrate.
This is explained in a paper on yeast alcohol dehydrogenase, but only the abstract is freely available. They claim to have evidence against Pauling’s idea, but I am not sure the extent to which this is generally accepted.

Paper can be found here.

Legally? I don't read Russian.

Sci-hub is quite a lot, but it is surely not legal.


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How did life survive in the Snowball Earth theory?
There is this theory that says that the whole Earth was once covered with snow and ice. How was it possible for life to survive this extreme state?

If you refer to ice ages, at that times not the complete earth was covered in ice. Do you have a source for this?

@Chris, the OP is referring to the Huronian glaciation, when the earth's surface became entirely frozen. We call this "snowball earth": https://en.wikipedia.org/wiki/Huronian_glaciation

@GerardoFurtado Interesting, I didn't knew that. Always something to learn here :-)

Indeed, always! :-) That's the beauty of studying sciences.

First, the "snowball earth", despite favoured by many scientists, is not a theory in the scientific meaning of the term, but a hypothesis, because it is not widely accepted: https://en.wikipedia.org/wiki/Snowball_Earth#Scientific_dispute. Some models show sea ice at the equator, while some more sophisticated climatic models failed to form sea ice to the equator.
But let's talk about the most famous "snowball earth" event, the Huronian Glaciation. It happened just after the "Great Oxygenation Event": the massive amounts of oxygen (O2) released in the atmosphere decreased methane concentration, and temperatures plummeted. It's supposed that Earth's surface became entirely or nearly entirely frozen.

How was it possible for life to survive this extreme state?

It's worth mentioning that, by that time (2 and half billion years ago), there were only unicellular organisms (and probably no eukaryotes, just bacteria). There were no pluricelular organisms nor life on land (outside water). Thus, having in mind that there were still a lot of liquid water (under the ice), it's easy to see those bacteria surviving. Sure, many species have become extinct, but not all life forms were threatened.
About the Huronian glaciation:

http://link.springer.com/referenceworkentry/10.1007%2F978-3-642-11274-4_742
http://www.sciencedirect.com/science/article/pii/S1674987113000297

The fact that there only existed unicellular organisms is indeed an important fact. Wasn´t flora life around?

2.5 billion years ago, only prokaryotes. If you call photosynthetic bacteria "flora", you could say that there was flora...


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How do you obtain specific mRNA transcript levels for comparison from a Hi-Seq dataset?
I have HiSeq data from mice exposed to two conditions. I would like to answer the following question: "Is there a significant difference in mRNA transcript levels when comparing condition A to condition B?"
To answer this question, do I have to map my HiSeq reads to the reference genome to determine expression values or is it acceptable to use a small set of reference sequences of interest (approx. 50)? These particular sequences are not annotated on the reference genome. What are the downsides to not mapping to the genome?

how many mice are in your groups? do I understand you correctly that you have 50 mice in your reference group?

Seven in each group of mice. I would like to examine transcript levels for 50 mRNA fasta sequences. Essentially, can I "fish" for only these 50 sequences out of my Illumina Hi-Seq data without mapping to the mouse genome first? Will a lack of mapping to the genome falsely inflate what is identified as one of my sequences of interest?

Does this help?  https://www.biostars.org/p/75462/  It's a post about aligning sequences using a reference genome without annotating.  Or this article about de novo transcriptome assembly: https://genomebiology.biomedcentral.com/articles/10.1186/s13059-014-0553-5

@VanceLAlbaugh Thank you for your thoughtful edit and clarifying questions

@GaiusAugustus The biostars post is helpful, thank you. Based on that post it seems that what I am proposing is the "quick and dirty way to do basic gene expression experiments", albeit perhaps less excusable given that there is a great deal of information known about my organism versus that of the biostars post. I feel that de nevo assembly will not be necessary for my purposes but thank you for the reference. I will proceed both with and without the genome mapping to compare.

It's going to be hard for you to generate an RPKM or RPM value without % aligning to the genome, or the transcriptome.  Your read counts will have no context.


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Does having to urinate imply that you are becoming dehydrated?
This question came up after I read another question that coffee does not dehydrate you, but instead acts as a mild diuretic.
Does having to pee imply dehydration?
Allow me to elaborate: When you have to pee, that liquid is already in your bladder, right? If so, it is not in use by cells. However, since that water must have been previously absorbed in the body in order to be filtered through the renal system, having to pee indicates that you are less hydrated than you once were.
So in the case of coffee, while it doesn't directly dehydrate you, doesn't it indirectly dehydrate you by causing you to pee?

Hm, aren't you forgetting that you are ADDING water to your body by drinking coffee?

Also, there are two common ways of describing "wanting to pee". There is "frequency" (feeling like you're going a lot) and "urgency" (if I don't go in the next 10 seconds my bladder is going to explode).

There are two parts to answer this question. "How does my body know when to pee? Do diuretics affect that, or my hydration?"

Does wanting to pee imply dehydration?

In summary: Not necessarily. Your body might have plenty of water, but the bladder is full. Alternatively your body might be dehydrated, but the caffeine and normal body functions have filled the bladder and you'll need to pee.
Caffeine as a diuretic.
Caffeine belongs to the Xanthines, and inhibits reabsorption of Na+ by cells, and increases flow rate of filtered fluid through the kidneys. This reduces water content in your body. In other words diuretics do dehydrate you, specifically caffeine will. Coffee and tea (very low doses of caffeine) are mild diuretics.
Bumetanide, Furosemide, and Torsemide are examples of stronger diuretics, and hydration should be monitored closely.
On the other hand, the colour of your urine may indicate your hydration.
The below image is a crop of this Cleveland Clinic infographic.

How does my body know when to pee?
The bladder is hollow and slightly elastic, so as it fills with urine it expands. A healthy bladder can hold up to 2 cups of urine comfortably for several hours. When the bladder approaches its maximum capacity, signals are sent to the brain, letting the body know that it is time to urinate.
But it's critically the kidneys that are controlling the production of urine.
Caffeine increases the production of urine from the kidneys, and fills the bladder quicker. However, the increase in production of urine from a cup of coffee is more than offset by the water in the coffee.


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Why are the positions of simple sequence repeats at the gene and related transcript sequences different?
I'm working on the simple sequence repeat (SSR). I found just one region (about 21 bp) with an SSR motif in a given gene. This gene has multiple transcripts, with the same SSR motif located in various positions of different transcripts. Could you please help me understand how this happened?

Because of Alternative Splicing, suppose an exon occurs neither the first nor the last, then the same exon will come at various positions in different transcripts of the same gene. Thus, if your SSR of interest exists in that exon, you will find the same SSR located in the various positions of different transcripts.


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What Species of Duck/Waterfowl is this
I snapped a picture of this beaut in Irvine, CA, but I do not believe it is in the list given in "Ducks at a Distance: A Waterfowl Identification Guide" by Bob Hines (http://www.gutenberg.org/ebooks/18884). What species of duck is this? I believe it has the shape of a duck, but the colors are unfamiliar to me.

Credit to @Sᴋᴜʟʟᴘᴇᴛʀᴏʟ for providing the link to the pdf

I think this is an Egyptian Goose (Alopochen aegyptiacus), see the image (First from the Wikipedia article, the second from here, interestingly taken in Irvine):

You can see the distinctive spot around the eye and also the colored feathers on the back. Also, this bird is too big to be a duck.


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How to define a 'trait' objectively?
I'm coming across an increasing number of papers that use trait-based approaches for risk assessment, like this one . This paper defines a trait as follows:

Traits are the physiological, morphological, and ecological attributes
  of species or other taxonomic entities, which describe their physical
  characteristics, ecological niche, and functional role within an
  ecosystem.

I've wondered about using this sort of approach during some analyses of invasiveness potential in vector insects before and it's likely to come up during a systematic literature review I'm coordinating, but I'm concerned that devising my own trait classification system is likely to introduce confirmation bias into a study. Is there either a recognised methodology for trait definition, or alternatively some sort of accepted list or classification of traits that I could use, to minimise the chances of this? Is there a generally-accepted method to correct for phylogenetic correlation?
I've tried Google but the results mostly appear to be about trait theory, which is an approach used in psychology to study human personality.

you might want to see this http://biology.stackexchange.com/questions/37143/definition-of-phenotype

I think you this paper is what you need. The many concepts of trait are discussed in the functional ecology background. But basically, a "trait" can be defined as

" 'Functional traits’ are defined as morpho-physiophenological
  traits which impact fitness indirectly via their effects on growth, reproduction and survival, the three components of individual performance." (Violle et al. 2007)

Which are not so different from the definition you posted.
Some organisms have an improved methodology to collect and measure traits, like plants. But this is not common to all organisms.
As for the method to correct for phylogenetic correlation, this is a whole different field called Phylogenetic Comparative Methods. To choose the method you need to know the kind of data you have and the question being asked.

That looks like exactly what I need for the definitions part of the question. Thanks!


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Optimal pH of protein buffer? Basic principles to adjust buffers according method and analysis
Protein buffers such as PBST, which is used in western blotting, are normally adjusted to pH 7.4. When I try to find why, I find some information about optimal pKa for protein stability. Im not sure I really understand this, because it seems as through regardless of protein - most laboratories just use PBST with a pH of 7.4.
I am a bit curious about this, and have a few questions

Why is pH 7.4 set as the standard for western blotting?
For an optimal western blots, should the pH always be above the pI of the protein?(since 7.4 seems a bit high)
Should the pH of a buffer be at the pI of a protein for extractions before SDS-pAGE, and while washing NC-membranes after blotting?
How should one calculate the optimal pH, or evaluate/think about this when working with a protein?

That's the average pH of extracellular fluid and cytoplasm but it can vary. Compartments such as lysosome and mitochondrial matrix would have a different pH. Functional studies with native proteins should employ pH buffers with right pH. During western blots, proteins are denatured and so their isoelectric points/functional pH do not matter. The pH in SDS-PAGE is adjusted according to isoelectric point of glycine which helps in stacking of the proteins.

Thanks! So basically all you would care about is the pI of glycine, which is why pH 7.4 is the optimal pH for western blotting.

Well.. it matters for SDS-PAGE.. Not for the blotting step. A specific pH may have differential effect. I have not really thought about that. Good question. I'll look up.

Thanks again, this is interesting and I think its important to get the basic understanding of how things work. I will add this reflection as a point in my question above!

The correct pH depends on the properties of the proteins that you are trying to exploit. These are a good deal different between extracting proteins from tissue and performing Western blots.
Extracting proteins means keeping them soluble in aqueous solution. Working at the pI of the protein might be the worst choice as the protein then has no net charge, typically reducing its water solubility. An extreme case would be the highly basic histone proteins with very high pI values but best extracted at low pH.
After proteins have been run on a denaturing gel and transferred to a membrane for Western blotting, they are already denatured and bound strongly to the membrane through hydrophobic interactions little affected by pH except perhaps at extremes. For the blotting, you want to exploit the interaction of epitopes on the protein with the antibodies against them. During antibody production, the interactions between the epitopes and the antibody-specific receptors on the antibody-producing cells occur at normal extracellular body pH, 7.4. Using that pH for the blotting best recapitulates the charge state of both the epitopes and of the antibodies as they existed when cells producing the best antibodies were selected.


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How do marine mammals and penguins maintain bone density?
After much searching on Google with no results, I decided to ask my question here.
If I'm not mistaken, swimmers' bone density is less than normal; at least for lifetime swimmers. Astronauts also lose bone mass from the lack of stress (from gravity) on their bones; it stands to reason that being in the water and thus being weightless, would lead to bone loss.
So my question is, how do marine mammals and penguins maintain their bone mass?

Wouldn't bones nearly completely wither away in a weightless environment, such as underwater?

FYI, some of the densest mammalian bones belong to cetaceans. For instance, blainville's beaked whale has a mesorostral with 2.6g/cm3. Sperm whales have a bulla with 2.16g/cm3. For comparisson, human bone is around 1.7g/cm3 (source: Bones and Cartilage: Developmental and Evolutionary Skeletal Biology). Also, penguin bones are considerable denser than other birds.

My comment was to OP.

Thanks for the input. Question though, why do lifetime human swimmers lose bone mass, shouldn't the muscular activity required for swimming keep the bones strong? What about the hormonal component?

See this article

Do you have a source for the bone loss in human swimmers? any vigorous activity puts stress on the bone, swimmers are not massless they still have to push hard against the water itself to swim. Also consider the alteration of an organism during its life is very different then evolutionary adaptation to an environment.

I wouldn't refer to it as "bone loss" but more as bone strength. Lifetime swimmers spend a lot of their time in the water. As in water yes they are almost "weightless" and their bone strength will decrease because it has gotten used to the "low" gravity. Now in the case of penguins and other Marine mammals, they have adapted to the force that the water applied to them.
Here is a link to a picture of penguin's flipper.

This answer gives two reasonable hypotheses for a putative answer, however it lacks reference. Could you please provide some evidence for your suggestions?

How do you define bone strength? What do you mean by "adapted" as in "in the case of penguins and other Marine mammals, they have adapted to the force that the water applied to them"? It says nothing about the bone strength, however it is defined, and its evolutionary adaptation. What does the picture say about bone strength of a penguin's flipper?


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Can it be said that proteins determine phenotypic traits?
I am not ignoring the function of genes, or the gene-environment interactions. What I want to know is if behind every "observable characteristic" we can find a protein -or a group of proteins- which is responsible for that trait. The idea comes from a school curriculum, and it is expressed as follows:

In this unit the core topic is the central role of proteins as executors of fundamental biological functions and thus as responsible of an organism phenotype...

Thank you in advance.

Yes proteins are executors of our fundamental biology. But the proteome is the critical word here (takes into account spacial and temporal features of all proteins). For example if you had a perfectly functional ion channel, but with low expression, then a different phenotype would be exhibited than a perfectly functional ion channel that was expressed in higher amounts. The proteins are the same but the proteomes are different.

A general note on phenotype
I think that the meaning of phenotype has become more complicated over time compared to what it was when it was first coined. As you said, in simple words, a phenotype is an observable characteristic of an organism. However, for Mendel, an observable characteristic would have been only an obviously gross change in the morphology whereas with the current state of technology we can observe the assembly of macromolecules inside a cell. We can also study the physiology of an organism/tissue/cell in real time using various sensors. We can also observe the gene expression profile using different technologies (with different levels of sensitivity and resolution). The question now is, can you call such a characteristic (for e.g. an assembly of some complex in a cell observed by cryo-EM) a phenotype? In other words "what is an observable"?

Coming back to your question:

Can it be said that proteins determine phenotypic traits?

Yes.

Are they the sole determinants of phenotypic traits?

No. A phenotype is usually determined by many factors. Many genes are involved, which may code for proteins as well as functional non-coding RNAs. However, if you make a comparison, proteins seem to be more versatile than ncRNAs (also most of them are involved in direct action). So you may say that proteins are the primary determinants of most phenotypic traits, but I would generally advise that such generalizations should be used cautiously. I'll give a counter example in favour of RNAs. If there is a mutation in a tRNA or a rRNA, you'll see a great effect on the phenotype.

About the question "what is an observable?", I would agree with this: "...the term phenotype includes traits or characteristics that can be made visible by some technical procedure." (https://en.wikipedia.org/wiki/Phenotype#Difficulties_in_definition)
In the case of an tRNA/rRNA mutation, that would lead to a change in the amino acid sequence, am I wrong?

@PiQ It may not necessarily cause a change in the amino acid sequence (it could increase the error rate). It might just reduce the rate of protein synthesis.

Yep, that is correct in general. However, sometimes proteins indirectly determine phenotype. For example, the skin colour is dependent on melanin (not a protein) but the melanin producing cells work because of proteins. Further, melanin production is also controlled by proteins. Thus, skin colour is not DIRECTLY determined by proteins but because proteins produce the pigment melanin, proteins are ultimately the reason behind our skin colour phenotype.

You might modify your answer to clarify your statements "…is controlled by proteins" and "proteins produce pigments". I don't think a novice would understand them. I would say that non-protein substances responsible for a phenotype are produced in reactions that are catalysed by protein enzymes. That's why proteins determine the phenotype.

When you say "in general", is it explained in the next sentence? Or were you referring to something else?


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How do afferent mechanoreceptors work on the finger pads?
I'm having some difficulty understanding how the afferent signals are sensed in the finger pads. My understanding is that for mechanoreceptors, as the indenting force increases, their effective response also increases. Additionally, as contact area increases, so does the overall stimulus area and hence a larger response amplitude from the mechanoreceptor afferents should be found.
However, the results shown in this article in the Journal of Neuroscience that discusses this physiology shows contradicting results. In this article, the closer the dot spacing (1 mm or less) and the higher the contact area the lesser the receptors seem to fire, and stop firing altogether at about 1 mm dot spacing?
Why would this occur?

I made some edits to your question... not trying to change your question but only clarify it for other users so you can get a good answer from someone.

Unfortunately, this is not a "please hold my hand and walk me through this paper/concept/idea" site. I strongly suggest you take the [tour] and carefully read through the [help] to learn more about the site, including what is on-topic and what is not, and how to ask a good question.

This is a really tough question to answer. I skimmed through the linked paper (authorative authors there!) and the last part of the discussion goes into more detail on this question, including references to other papers. I may get back to this question, but it's going to take me a while. Good question! +1

Can you also pinpoint the figure numbers so that the actual issue is easier to spot?

This took me a while and my apologies - I have to add that it's an excellent article with a set of awesome authors and it was an enlightening read. Thanks so much - I learned a great deal :) I'm actually writing a paper on this stuff as we speak, so questions like this make this site so fruitful :-)

Short answer
Tactile grating stimuli with a higher line density (number of lines per surface area) and applied with an equal force result in a lower amount of pressure per unit of skin area under the grating, and hence smaller afferent responses.
Background
In the Discussion section of the linked article (Phillips et al, 1993) the authors explain on p. 838 (left column) that the reduction in the response of SA afferents (slowly adapting fibers) when grating density is increased is due to the higher proximity of neighboring areas of stimulation, i.e., the lines in the grating stimuli are more closely spaced. The authors reason that this leads to a reduction in the effective stimulus (i.e., compressive strain) at the receptor terminal due to the presence of adjacent stimulus elements that distributes, and hence reduces the skin loads per line.
Reference
- Phillips et al, J Neurosci; 12(3): 927-39


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Sensory neurons, interneurons and motor neurons - recognizing the type of neuron at microscopy images
Is there a simple way to distinguish between sensory neurons and interneurons and motor neurons at the microscopic images?
I read a little bit about Open Connectome Project. Is there a possibility of automatic (computer - algorithmic) recognition of the aforementioned types of neurons?

Classifications are based on shape, number of connections, projections of axon and such. Thus there is definitely a possibility of automatic recognition if its not already being done.


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Are all prokaryotic promotor regions the same, or do they differ depending on the product of the transcript?
Are all prokaryotic promotor regions the same or do they differ depending on the protein encoded by the mRNA being transcribed?
What I am thinking is that if activator proteins are a method of transcription regulation, then shouldn't promoter regions differ so that activator proteins have an effect on the  expression of a specific gene?
Please tell me if this is correct, or if I misunderstand something.

Sorry, english is not my first language....

Definitely not. There is a ton of variation from gene to gene, otherwise, as you say, regulation wouldn't work.
That said, the word "promoter" sometimes gets used in different ways. Especially in eukaryotic systems, people will sometimes say "promoter" to mean the entire region upstream of the gene where transcription factors can bind (for yeast, typically within a few hundred base pairs, for mammals, could be many kb away). These regions are extremely variable from gene to gene in all organisms. But for bacteria, the word is more commonly used in a much stricter sense, to mean the region where the polymerase and the associated sigma factor binds -- this is typically one or two very short stretches of DNA (maybe 5-10 base pairs) very precisely located at a specific position from the start of the gene. These regions are much more conserved, but there will still be some variation. Check out http://nar.oxfordjournals.org/content/40/7/2907.full for a very detailed characterization of some of these sequences in E. coli.

This is not my area, but as nobody has answered yet I'll start the ball rolling to try to encourage a more authoratative response.
Long, long ago, before the internet, when I was a post-doc and the emphasis was on bacterial rather than eukaryotic gene expression, I remember hearing something about specific sigma factors differentially controlling initiation of different bacterial genes. A quick internet search brought up something in Wikipedia (section on Specialized sigma factors), a recent review on sigma factors by Chandrangsu et al. (2014), and a more general review of the regulation of bacterial transcription initiation by Browning and Busby(2004).
In general it would appear that you are correct in thinking that promotor regions will differ to some extent, depending on how they are regulated.


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Is brachydactyly due to mutation?
I have this so called "clubbed thumbs"also known as brachydactyly.
It is of D-type.
I searched for it on internet and found that it is a dominant inherited disease.
But to my surprise,none of my family members have such thumbs.Not even  my great grandfather/mother had this type of thumbs.
If it is a dominant trait then why didn't it show in any other family person. Is it that I have had any mutation in my genes?

You are correct that brachydactyly is a dominantly inherited disorder and is usually caused by mutations in the BMPR1B gene.
However, not everyone who has the mutation, will have clubbed thumbs (the phenotype). This is due to a phenomenon known as penetrance and expressivity.
Penetrance essentially is an all-or-none phenomenon whereby certain individuals who have a mutation express the phenotype (in this case, clubbed thumbs) and some do not express it at all.
Expressivity is when individuals who have the mutation, will express the phenotype to varying extents. For example, only one thumb may be clubbed.
I hope this helped.


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Found an unknown species, whom should I contact?
My father has found a very queer looking insect. He'd been watching it for over half a year on the tree right next to our house  and in all this time hasn't been able to find anything like it on the internet. Yesterday it died, and we found it on our porch, dad's put it in an air-isolated box for identification, please let me know if you are interested. We can send a picture of it for you via email or any other provided way, or please let us know what we should do it and whom we should contact. We live in Sydney.
Thank you, best of luck in everything!

Welcome to Biology Stack Exchange. Species ID questions are quite common on this site - try uploading some photos of the insect and adding them to your question, and there is a very good chance that someone here can ID it for you. Make sure you also add the species-id tag. For insects I would advise adding details of the weather when it was seen.

And just a side note. Don't be afraid to actually post the picture. It isn't easy for someone to just steal it and claim it as his own. Plus it would be very easy to prove that you were the original poster. So I would recommend to just post it here. You will get more feedback that way.

To extend on Ria's point, even if someone stole the image, they can't steal the insect's corpse over the internet. If it really is a new species, that corpse will be more important evidence than any photograph.

As above, please upload a picture of the specie you've found to confirm your recent findings. It would be a good idea to contact a professor at a university in Sydney that works in the relevant field as he would be known as a reliable source for further referrals

It is kind of assumed that if you know enough t actually identify something as new you will know what to do about it. Searching the internet is not a great way to identify an organism.

Has it been identified yet?

Natural History Museum; UK, is a good resource.
http://www.nhm.ac.uk/take-part/identify-nature.html

According to their webpage;
Have you found a strange bug or a new plant in your garden, or stumbled across an intriguing animal bone or fossil while out on a country walk? Let us help you find out about it.
Four ways to identify your specimens and sightings

Check our selection of ID guides and apps below to see if one is relevant.
Upload your photos and descriptions to our online identification forums for help from our scientists and other naturalists. Please tell us as much about your specimen as possible, including a detailed description, where and when you saw it, and its approximate size.
Post your specimen or photographs to:
  Identification and Advisory Service
  Natural History Museum
  Cromwell Road
  London SW7 5BD
Visit the Centre for UK Biodiversity at the Museum and show your wildlife or geology specimen to a member of our ID team. Drop in between 10.00-12.00 and 14.00-16.00:
  Monday to Friday (except bank holidays)
  the first Saturday of the month (or the second Saturday after a bank holiday weekend)
  If you need to discuss the best way for us to help with your enquiry, call our Identification and Advisory Service team on +44 (0)20 7942 5045 or email us.

See more at: http://www.nhm.ac.uk/take-part/identify-nature.html#sthash.UGsoIYvE.dpuf

They also have provided some downloadable apps and pdf document resources in their webpage.

Although the UK does have a history of collecting everyone else's specimens, whether they were solicited to do so or not, I would suggest not sending anything (live or dead) across international boundaries, in this context :)


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How many Amperes to kill fruit flies and mosquitoes?
As above. Making a homemade trap to kill them all.

If you were to presume that the resistance of the arthropod is about 1000 ohms, then it'd require about 2 amperes.

I have this bug-zapper in the form of a tennisracket. It takes only two AA batteries of 1,5 Volt. It is succesfull in killing mosquitoes but often fails at big flies.

It's not the Amps that matter, it's the voltage. humans are safe up to about 80V DC. 4V is like moving energy in a slow train, 1000v is like moving energy in a bullet. Therefore, 1000V also can arc more easily through the air. If you use 240V at 0.1 amps, that's 24W it's probably quite enough to kill a mosquito. A 200W source in 4V will not hurt a human, but in 3000V it will kill him easily. I can't confirm that 20W at 300V is not dangerous to a human. You will probably find yourself using an CC Constant Current supplying about 10W at a low voltage and upping the voltage a bit to zap the moz.

Current kills. Voltage is potential difference.

I am not an electrician and may not be the right person to answer this. But I recently read this paper (Keller et al., 2016) and maybe this helps ... Keller et al. (2016) have a look at laser induced mortality in mosquitoes. It might not be fully adaptable to your needs (there is not much context in your question, though; I have no idea what kind of trap you are trying to build ... if it's going to be a fancy laser trap, the cited paper will be your perfect guide through the building process - however, you have to figure out the automatic camera detection (they are planning that, too) for yourself as they have not published that, yet).
Now, giving a concrete answer: Keller et al., (2016) found that lasers with 0.5W to 4W are sufficient for efficient mosquito inactivation. If you are planning to use a plug bucket (230V) this would be (0.5W/230V=) 0.002A to (4W/230V=) 0.017A. You could also set this up with a 9V battery. With a 4W laser this would be (4W/9V=) 0.44A. Using a lithium-based battery this would last for about (1000mAh/(4W/9V=0.44A)=1Ah/0.44A=) 2.27h total laser time.
I do not know if this easily translates to electric current, though ...

What did Keller et. al. use as the ohms of the insects in their investigation?

They did not assume any. They tried this with a variety of powers and pulse lengths and then measured mortality rates.

Sorry, laser's not applicable. The trap's made up of loads of exposed wires alternating between live and ground over a food source, waiting for a fly/mosquito to close the circuit to (hopefully) fry them. Not going to plug it into the mains, though. Too dangerous. Thanks for the power details.

@user3635998: Ah, ok. Maybe you should have a look at how these  electric fly flaps work, I think they are powered with two AA batteries and get up to 1000V with a voltage converter.

there's an electronics topic for that, it states: Most flyswatters conform to electrical safety standards for humans: a limit on the charge stored in the capacitor. A discharge of less than 45 µC is considered safe, even in the unlikely scenario that the current from a flyswatter would be flowing from one arm to the other arm, partly through the heart.1 This means that the capacitor of a 1000 V flyswatter should be less than 45 nF. Due to this precaution for humans the initial shock is usually inadequate to kill flies. a limit on the current after the initial discharge. The maximal continuous current of most flyswatters is less than 5 mA. This current is safe, even when flowing from one arm to the other arm
here: https://electronics.stackexchange.com/questions/19367/can-i-test-bug-zappers-voltage-with-multimeter
there is a slow mo zapper here. https://www.youtube.com/watch?v=oUJLgKkmllA


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What does completing the course of antibiotics accomplish?
This article from the WHO states:

If you stop treatment early, there is a risk the antibiotics won’t have killed all the bacteria that made you sick and that it will mutate and become resistant. This will not happen to everyone – the problem is that we don’t know who can safely stop treatment early.
By taking the full course prescribed by your doctor, even if you start to feel better earlier, you increase the chances of killing all of the bacteria and reduce the risk of resistance.

A clarification: When the WHO says “[bacteria] will mutate and become resistant,” they are implying that using antibiotics will create a selective pressure for bacteria with antibiotic resistance, arising most likely through mutations, correct? Bacteria don’t mutate into antibiotic resistant strains in direct response to a drug; rather, antibiotic resistance is a phenomenon of natural selection. I’ve seen so many articles about antibiotic resistance worded this way and I feel it’s simply wrong.
Presuming I am correct about my first point, here’s my true question: Say you’re prescribed a course of antibiotics. It kills the bacteria that are not resistant to the drug and leaves those that have developed resistance behind. What does it matter if you finish the whole course? Taking more of the drug is not going to affect the resistant strain. The only rationale I can think of is that it would kill off any residual non-resistant bacteria and prevent them from gaining resistance (via conjugation) from the resistant kind. However, is this the only mechanism whereby completing the course prevents the rise of antibiotic resistance? How effective is this at all at preventing the rise of antibiotic resistance in the bacterial population in general (i.e. beyond the host)?

Good points. Yes, it is worded misleadingly. Resistant bacteria are selected for in the presence of antibiotic. when you stop an antibiotic, levels are still present for a while and it may select for more weakly resistant bacteria (that can survive at low levels of the antibiotic - very common.) Stopping your antibiotic early mainly means you've left a lot of bacteria behind (that colonized part of your body) and reinfection is possible, requiring yet another course of antibiotics. It's the "yet another course" that confers the increased possibility of selection of a resistant strain.

"it will mutate" could also be read where "it" is the population of bacteria and not the individual bacterium.

As anongoodnurse has mentioned in the comments, an incomplete course can allow weakly resistant stains to expand their population. As you mentioned, the antibiotic does not cause the bacteria to mutate but it kills all strains that do not carry the mutation that provides the resistance. This mutation arises randomly (in certain cases it can be acquired by other mechanisms such as lateral gene transfer).
Many antibiotics may not kill the resistant strains but may still impose some burden on their growth. Now, with a complete course you would have eradicated a lot of strains and only a few resistant strains would be left. The probability of extinction/eradication depends on how many individuals are present in a population. When you do a complete course, then the chances of extinction of the resistant strains due to random birth-death process (and attack by the immune system) would be high because of their low numbers. However, with an incomplete course these bacteria would quickly expand their population once the antibiotic is gone. Now, with the next round of antibiotic treatment you would have more number of resistant cells to kill. As again pointed out by anongoodnurse, this will lead to further selection of resistant strains.
There are additional factors:

Microbial community (colony) is more than just a bunch of cells. They can protect each other.
There are some cells called persisters which are not really genetically resistant strains but tolerate the drug because of adaptive phenotypes. Again, the microbial community plays some role here.

Furthermore, resistance comes in many strengths. A weakly resistant strain is less likely to die from an incomplete course of antibiotics. But a weakly resistant strain is many more times likely to become a strongly resistant strain...

@Aron "a weakly resistant strain is many more times likely to become a strongly resistant strain” Why is that so?

@lightweaver Imagine one strain randomly mutates an enzyme that breaks down the antibiotic. But it's a very weakly binding enzyme. Then imagine how much it would take to optimise the enzyme.

@Aron Could you give an example? Maybe it’s the lack of experience but I’m not quite imagining it.

@lightweaver An enzyme that can break down the antibiotic could requires thousands of mutations. But it might only be good enough to break down half the antibiotic in your blood. So if you take a full course, the bacteria that makes it still dies. However, if it survives a half course...all its rivals are dead. It multiplies, and all the bacteria has a weak immunity. All of them mutates...maybe 10 or so mutations. But out of the trillions of bacteria ONE hits the jack pot, and evolves the 10 mutations required for FULL strong immunity. Congrats, you just killed man kind.

You also have bacteria that go through dormant phases, they are not resistant they just are not active so they are unaffected , following the full course will catch these guys when they wake up. Otherwise you are just instantly reinfected when they become active again. TB is notorious for this.

@John those cells are called persisters (fancy term)


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PBST vs. TBST buffer in western blotting
What is the advantages and disadvantages of using either PBST or TBST in western blotting, or while working with proteins in general? Are there other buffers which are also used for western blotting, or washing steps while working with proteins?

In my experience it actually doesn't matter which of these buffers are used for washing. I think this is much more a flavor of the respective lab.

I used to work for a company well-known for its modification state-specific antibodies, including phospho-specific ones, and they actually performed extensive in-house testing of PBST vs. TBST in Western blotting. Part of the reason the company chose to recommend the use of Tris-buffered saline over phosphate-buffered saline based buffers was the clearly-demonstrated fact that some phospho-specific antibodies just didn't work as well in PBST as they did in TBST, for whatever reason. As I recall (it's been quite a while since the testing was performed), some of the antibodies tested gave weaker signals, with more background, and one in particular just didn't work at all in PBST - no signal whatsoever. For that reason, the company chose to standardize its recommended Western blotting protocol with TBST, and it's been that way ever since.
That being said, I'm sure that the majority of antibodies out there would work just fine in either buffer. However, to be on the safe side, if a company states one buffer or other in their protocol, it would be best to use the recommended one.


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Natural rulers: What organisms have the most regular size?
What organisms exhibit the most precise length scales?  E.g. are there trees with rings that are of very precise thickness, or single celled organisms which are all very nearly the same length?
In particular, it seems to me that most lengths in nature are precise only to about the 10% level.  Are there any organisms which exhibit length scales considerably more precise than this?
(Bonus points if you can tell me the most precise mass scales in nature, too.)
PS: This question is partially motivated by ongoing studies by physicists as to the origin of length scales in biology.  See this paper for an example.

You mean to ask which organisms have least variability in their sizes (and converse)?

@WYSIWYG Essentially that's what I'm asking, with the caveat that I'm not necessarily concerned with overall size, but rather the size of any component of the organism.  So e.g. in the case of (say) trees, the overall height of a tree need not have small variability, so long as the growth rings have very consistent size from one tree to another.


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Does my brain have limited learning capacity or memory?
Lately in my free time I like to read books, articles etc etc.. What I have discovered, or maybe it is an illusion(lack of concentration or something), is that I usually forget most of the material that I read about, and have lost my ability to remember names of objects, planets or whatever I read about. Does that mean that I am over-reading or that my brain has limited memory/learning capacity? I should also mention that I have an IT job, which you know requires high concentration throughout the day.

This sounds more like a health advice. Please edit your question to clearly outline the scientific premise and add details and examples if needed.

this is a much better fit for the https://psychology.stackexchange.com/

I disagree with @John comment, neuroscience of the memory is one of the areas of biology.

technically all of psychology falls under biology, but we have a stack devoted to that one subject that will be better able to answer.

The Bekenstein bound limits the amount of information that can be stored in any finite amount of space, so a fortiori your brain does have a limit. Of course, the limit imposed by this bound is vastly higher than the practical limit (if it weren't, our brains would become black holes).

Short answer is yes, the brain has a limited capacity. It's a large but limited physical system with numerous but countable connection, it follows that it must have a limited capacity.
Learning wise, we usually say we learned something when we can recall it at will. I'm not sure I agree with that definition, but if this were the case, learning would be intimately related with memory capacity, so the answer would also have to be yes.
All this without taking into account energy consumption and metabolism which would impose also further limitations, e.g. I bet you that if you switch the activities around and start reading in the morning and try to work at night your work would suffer but your retention capability would improve (just a guess). Now, is all this relevant? I can't say, we (humans) might be working already at full capacity, in this case it would be relevant, or we could be very far away from this theoretical limit and all this is moot.
If that was really your question then you're done.
However, you seem to be preoccupied with a mismatch between the expectations of your own retrieval abilities after a particular "type" of activity and what you can actually remember.
I would be remiss in I didn't say that I agree with WYSIWYG's comment, I'm sure this is not the place for health advise. So with that in mind my 2c:
your expectations can be misaligned, that's always a possibility; your retrieval abilities can be impaired, which means that you learned fine but cannot recover it; or may favorite: you might just be tired! IANTTOAD


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"X" in stock solutions
We prepare buffer solutions with concentration in terms of x, for example 50x TAE buffer. How do we come up with x? Is x related to molarity in some sense? And how much can we concentrate the solution. We have 50x TAE but usually 10x TBE buffer, why is it so?

X simply stands for "times" like 50 times concentrated. And there are limits of how much of a certain substance can be dissolved in water. This limits stock concentrations.

Thanks for the comment @Chris. If X stands for "times" as you say, how much is 1 time, i.e, 1x, is it 1M, 0.05M as given here link? And is it same for all solutions?

That depends on your final concentration. If it is 100mM, then 10x is 1M (1000mM), 100x would be 10M.

1x is the final working concentration of the solution (it could be anything depending on the type of the solution). Stock solutions are made at a higher concentration; if it is 10 times more concentrated than the working solution then it is 10x.

Why 50x TAE but 10x TBE?

The salts in TBE precipitate at higher concentrations. In other words, the salts in TBE have lower solubility compared to those of TAE. Therefore stock solutions of TBE are usually 10x or even 5x.

Thanks @WYSIWYG. Can you tell me how do we decide what should be 1x concentration by citing few examples?

@Biginer How to decide the 1x concentration: the first time when someone does that, then they 1. use rational guess 2. use trial and error, and settle on a concentration that is most optimal for the task. Subsequently, others follow it and tweak it when necessary.

Thanks @WYSIWYG. I came to the same conclusion when I discussed it with my peers. Thanks a lot!


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How many co-expressed genes would be expected in a tissue?
I am working with gene expression microarrays of tumor tissues and I want to use a program to find the clusters of co-expressed genes in order to know if some particular genes are co-expressed with another genes and which genes are those.
As I must do this for many microarray experiments and I have read that there are a lot of genes that have constant expression in a tissue, I ranked the gene by its variability (coefficient of variation) that they had in the experiments and keep the first 3000 genes to search their co-espression. Then I employed a program (a bioconductor package) to find co-expression.
Now, I was expecting to find several clusters of genes (no particular reason for this) in the experiment that I was using as example, but instead of that, I only find one cluster of co-expressed genes of about one hundred, and the others genes didn't were co-expressed.
My question is: this result could have a biological sense, or I'm making a terrible mistake in some place?, Thanks in advance.

You clustered the genes based on what?

It's really not possible to interpret your analysis when we don't know what the data is. How many samples (array hybridizations) are there? What are the conditions, do you expect a good diversity of expression profiles? Is there any indication that the data is good quality, like concordant replicates? Many publicly available data sets are small / poor quality and cannot be used for cluster analysis.

Also, you might want to check out Olga Troyanskaya's lab at Princeton, http://reducio.princeton.edu/cm/ogt  They have worked on large-scale coexpression problems for some time and provide software libraries that solve many of the basic problems.

Check the beautiful publication of Daniel Ramsköld et al. 2009, which holds the numbers for generally anticipated co-expression.
The specific level of co-expression, which applies to your scenario, will depend upon your tissue, your thresholds, and your definition of co-expression.
It you look for a co-change of some genes across different specimen (rather than co-expression), the number of genes that should change will depend upon the underlying biology (and thus prevent a general answer without considering the specifics of your specimen).

There are certain very well defined groups of genes that you would expect to be co-expressed, e.g. ribosomal proteins, proteosome subunits, splicesome components, VHATPase subunits; and each of these groups are co-expressed in different circumstances. It therefore looks to me that either your different tumour tissues (if that is the situation) are in similar conditions so you can’t differentiate these clusters, or that there is some problem with your analysis.


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Accommodation mechanism.
Can anyone explain how contraction of the ciliary muscles causes relaxation of the zonules? Please explain it anatomically i.e. the attachments of ciliary muscles and its relation with the accommodation process.

If you explain in more detail what confuses you and why (i.e. show us you've researched this and tried to answer it on your own), I'm sure it would have a better chance at attracting an answer.

When we study the attachment  of ciliary muscle, it appears to produce more tension on the ciliary zonules (& thus also on the capsules of the lens)-thus should cause flattening of the lens(i.e lesser convexity or more redius of curvature). But according to Von Helmholtz's capsular theory on accommodation, contraction of ciliary muscle causes releases of the zonular tension on lens capsule. My point is..... Ciliary muscle surrounds and holds the lens capsule, so when it contracts it should shorten in length....thus causing more tension on lens capsule(which is clearly against the theorem)

I searched medical books on anatomy and physiology and different medical websites for it. I also tried to find an explanation in YouTube but found no video satisfactorily explaining it. So I posted it here.

I think the problem you're having is in the way you're thinking that all contracted muscle shortens, and all relaxed muscle lengthens. But the cilia is different. The ciliary muscle is a circular or radial muscle (like the orbicularis oris of the mouth.) Thinking of it like a sphincter muscle is much more helpful.
The ciliary muscle is attached at its periphery to the lens by suspensory ligaments (aka the zonules). Think of the the lens, the suspensory ligaments, and the ciliary muscle as a single unit with only two components whose length can change dramatically: the cilia and the lens. The length of the suspensory ligaments is more or less fixed.
This may be counterintuitive, but for seeing at a distance, the ciliary muscle relaxes, and the muscle in effect becomes "thinner" (think of the capitol letter O in a thin font, or think of a shpincter muscle; when the latter relaxes, it becomes thinner and more spread out). This means the "decreased distance" taken up by the ciliary muscle must be made up for by the suspensory ligaments and the lens: The tension on the suspensory ligaments increases, causing the lens to assume a flattened shape.
For near vision, the opposite occurs; The ciliary muscle contracts (think of the letter O now very bolded - it takes up more of the fixed distance.) This allows the ligaments to relax, allowing the lens to assume a more rounded shape.
Does that make any sense at all? I hope so, because it is essentially how it happens.
I did find a YouTube video; unfortunately it explains it pretty much like I did varying in a few details.
Morphological Study of the Accommodative Apparatus in the Monkey Eye Way way complicated; it's better to just believe me.


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What is the cause of the packaging capacity limit of recombinant adeno-associated viral (AAV) vectors?
I am working with recombinant AAV and I am puzzled by the limit of their  packaging capacity (around 5Kb ssDNA), that is really a huge drawback for gene delivery.
I have tried to look more in deep into this problem but I couldn't find any paper that could clarify what is the reason of this limitation. Is it the size of the capsid or something else? I suppose that it is the size of the capsid but there are smaller capsids able to accommodate more DNA so it could be due to other factors. Either way, I could not find the proper literature to solve my doubts.
Do you know the answer? Or, can you point me to good papers on this topic?

Cf. http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2835018/ and http://www.ncbi.nlm.nih.gov/pubmed/17328683

It's not the size that counts, it's how you pack it (sorry couldn't help myself).
Wild-type bidensoviruses do a lot of interesting things to pack genome like splitting its genome into two virions(1), inverted terminal repeats (ITRs) that form panhandles that can stack instead of hairpins (2), and overlapping positive and negative ssDNA strand ends (2 and 3) that clip closely together.  So on the whole, I don't know that it's a fair comparison.
Also remember that you can go too small with AAV genome size, not just too large (if you go too small you will even get 2 genomes per virion).
If you want to orderly pack ssDNA (or DNA in general), you have to use some nucleoproteins to help.  Free ssDNA has the added problem of being unstable.  Going back to basic biology, you can think of nucleosomes allowing the packing of eukaryotic DNA at a much higher density than would otherwise be energetically favorable.
Viruses use a wide variety of methods to pack genetic material, one of which is through binding directly with the capsid as with AAVs.  One should remember that the capsid proteins also are tasked with entry into the cell, trafficking, and entry into the nucleus.  All while protecting and packaging the genetic payload.
AAVs, being small (wt is ~4.7kb w/7 proteins), only have the capsid proteins VP1-3 to work with.  The ITRs are known to be the cis active sequences of the genome (4, 5), and these ITRs form normal hairpins.
A portion of the AAV ITR binds (hooks onto) an interior facing portion of the VP3 gene (6, 7).  But the virus has to push in this ssDNA, and it uses the Rep proteins to do it (8).  The Rep proteins (Rep52 and Rep78 specifically), are larger than the capsid proteins, and bind the ITRs at multiple sites.
While the exact insertion pore hasn't been identified at the structural level, the way I like to picture it is a series of hairpins have to bind on spokes of the Rep proteins to be pushed into the virion, where in turn the hairpins have to hook onto the inside of the capsid.  If you put too much ssDNA between your ITRs, then you lose the ability to be passed along (or bind) to both proteins in packaging.  You start to lose the tight packing onto VP3 found in wt virus, and instead get more sporadic VP3 binding (every 3rd as opposed to every other).
It would be incredibly hard to just "increase" the size of the capsid without messing up the assembly and the other functions of the capsid.  The VPs only oligomerize in the cytoplasm (9) suggesting that host proteins are required for proper assembly.  I think the process of successfully increasing the optimal genome size would be quite a task and achievement if done.  One would likely have to optimize additional ITRs, the internal binding bind sites on VP3, and the structure of Rep52 before such a thing would be possible.  Alternatively bringing in a novel or chimeric packing mechanism, but again this would be exceedingly difficult (probably prize worthy).


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How does gene editing in one cell affect other cells?
I have read about gene editing and I have some questions concerning this technology:

With gene editing, are we correcting the genes in all the cells or a single cell?
If the gene editing is done for only one cell, then how (or can) it affect other genes in other cells?

Gene editing is not done on a single cell but a bunch of cells. However, it still does not affect all the cells. Gene editing/therapy for the entire organism works in these cases:

Stem cells are edited, which give rise to many other somatic cells. Usually done in case of gene therapy in which the stem cells are obtained from the patient, edited and re-implanted.
Some embryonic cells are edited and reimplanted in the developing embryo which gives rise to what is known as a chimeric organism (some cells in the body carry the edit while others don't). Now some of the edited cells will give rise to germ cells which in turn give rise to gametes. In the next round of breeding the chimeric parents would give rise to a fully transgenic (with edited gene) organism.
For unicellular organisms, it is pretty obvious: you just use a selection marker or extensive screening to select for the cells carrying the edited gene.


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Is DNA sequencing coverage a function of sample purity?
How is coverage affected by the purity of a sample? And can coverage for a sample be affected by other things, like the library preparation or manner in which the sample was stored?

Coverage is basically affected because of concentration of the strands in the library. It can be affected at any of the several steps of DNA isolation, library preparation and sequencing.

So factors which go into coverage in short-read sequencing (and Sanger sequencing).

Depth is the obvious one. All else being equal you get a bang-on Poisson distribution of coverage over a genome, which is why more sequencing = more coverage.
The human genome and transcriptome is not all equal. Illumina sequencing is ultimately (most protocols) dependent on PCR amplification which means that any factors which influence PCR-amplification ability will also effect sequencing depth. The two obvious ones in this respect are: 1) Regions of extreme GC content and 2) Regions containing secondary structures (RNA mainly).
Alignment is the next major source of error for regions being 'dropped off' depending on how your aligner deals with multi-mapping reads. Some genomes have very young transposable elements which are highly similar to one another and very abundant. For most aligners that means that you can get either huge coverage or really small coverage depending on how the genome is assembled.
For low-input sequencing techniques you get major sources of DNA/RNA contamination from the enzymes, reagents and preparation of the samples. This can largely be minimized by having 'clean' samples and using large amounts of input DNA/RNA.

Depending on the sequences you're studying read-depth can be the only factor you consider or if you have difficult regions or specific questions you can deal with a whole swath of problems.
So in answer to your question: yes DNA quality will directly effect coverage. Sometimes you can 'just sequence more' to get over this problem, sometimes you need to go back to the drawing board.

The largest contributor to sequencing coverage (on a Hiseq, I assume) is how many reads you get, which mostly depends on what percentage of a flow cell you give it.  Obviously the rest that you mentioned contributes, and purity might be a big issue if you are looking for a subset of cells you were trying to isolate out of a larger pool, but the easiest way to get more coverage is to hog more real estate on the flow cell.

I don't understand what you mean by flow cell percentage - would you please briefly elaborate?

If you are not talking about short read technology, then it's more applicable.  Obviously, the most direct thing contributing to coverage is how many reads you have.  More reads, more coverage.


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Reason for variation in the site of onset of edema
What is the reason for the observed clinical difference in the earliest site of onset of edema in cases of different etiologies?
For example, in Congestive Heart Failure, it appears initially as pedal edema (edema of the dependent portions, in general) but in nephrotic syndrome (or other kidney causes) appears first as periorbital edema?
In both these cases, since the onset of edema is distal to the organ of the original pathology, it must be a systemic hematological effect caused secondary to the pathology that causes this specific pattern (as opposed to a localized portal hypertension in liver disorders initially appearing as ascites). Although all of these may progress to anasarca, what determines the specific pattern observed?

We should first recall how plasma (intravascular) fluids moves to the extravascular space. It can be approximated by Starling equation. Briefly:

The rate at which fluid is filtered across vascular endothelium (transendothelial filtration) is determined by the sum of two outward forces: capillary pressure ($P_c$) and interstitial protein osmotic pressure ($\pi _{i}$), and two absorptive forces: plasma protein osmotic pressure ($\pi _{p}$) and interstitial pressure ($P_i$).

See more: Starling equation
Primary edema localisation depends on the underlying pathophysiological mechanism that disturb the balance between aforementioned forces pushing fluid outside the vessels. Whether the Starling forces are alerted globally or locally, how long the edema develops etc.
Example from Harisson's Principles of Internal Medicine (I recommend the whole section regarding the edema):

Edema due to an increase in capillary pressure may result from an elevation of venous pressure caused by obstruction to venous and/ or lymphatic drainage. An increase in capillary pressure may be generalized, as occurs in heart failure, or it may be localized to one extremity when venous pressure is elevated due to unilateral thrombophlebitis.

In Congestive Heart Failure edema usually appears in places, where intravascular hydrostatic pressure is the highest. When the individual walks it is typically seen as ankle swelling, when the individual lies down, it is lumbar region edema.
While there is a decrease in osmotic pressure (like in nephrotic syndrome), the fluid moves toward extracellular tissues of relatively "loose"  structure (so for example periorbital region).
What is important, in some medical condition (eg. renal failure) different mechanisms may coexists and change the clinical picture.

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