text:
Sequence evolution simulation tool
I'm looking for a tool to simulate sequence evolution given a specific mutation model and birth-death model. I'm aware of tools and packages like INDELible, Seq-Gen and PhyloSim, but they simulate evolution along phylogenetic trees. What I want is to give a sequence and ask the tool to simulate the evolution of this sequence through N generations. The output should be a big bunch of descendant sequences. I've been thinking about writing such a tool myself, but it's always better to look for something already coded.

There exists a bunch of population genetics forward and backward (coalescence) simulation platforms. Here is a non-exhaustive list. They all differ and you'll have to go through their manual to see what is more adapted to your needs.
List of softwares
Here is a long list of such platforms. The list might arguably be a little bit outdated today though and many of these softwares might be slow or abandoned by their authors.
Most common softwares to my experience
Some are more known than others. Personally, I already saw uses of the following platforms in publication: SimCoal, Nemo, Slim and SFS_Code.
SimBit
Of course, I must indicate my own simulation platform SimBit. To my experience, SimBit is typically faster than Nemo. It is slower than SFS_Code and SLim for very small simulations but become much much faster than SFS_Code and SLim when you need simulations with a descent amount of genetic diversity.
Most important point about these softwares
You should compare softwares based on

Availability of the author to advice and bring new features

I won't talk about my personal experience here by respect for the authors!

Speed / RAM usage

I think SimBit is typically faster (and use less RAM) than Nemo. SFS_code and SLim are very fast for simulations with low genetic diversity but very slow for simulations with high genetic diversity.

Flexibility

Nemo, SLim and SimBit are very flexible (but they do different things) but SFS_code is not really.

User interface

I find Nemo error report pretty poor. I like SimBit error report. SLim interface is very nice (using eidos) and it even comes with a GUI (I have never used the GUI myself).

Free of Bugs

Several people have found important bugs in SFS_code (personal communication and experience).

Slightly more detailed comparison between Nemo, SLim, SFS_code and SimBit
I have personally used NEMO, SFS_CODE and SLim in the past (I now use SimBit only). So I can only talk about these 4 below.
All four models are well updated and maintained by their authors and of course everyone has a access to the source code.
SFS_code and SLim are very fast for small simulations. The issue of SFS_code and SLim is that the RAM usage and run time are exponential function of the genetic diversity. This means that some simulations can quickly become totally unmanageable. But it is a great tool if you expect little genetic diversity.
SimBit is typically faster than Nemo, at least for large genomes. This difference becomes particularly obvious for large genomes. One might think "Oh I am fine to wait a few extra days to get my results" but the difference in speed between two softwares can also be a difference in whether you have to wait a week or 20 years to get your results, so do not neglect the importance of running time and RAM usage without estimating your needs.
Nemo is very flexible when it comes to the order of life-cycle events (is migration before or after selection for example) and the newest version allows for simulating age structured population. SimBit is very flexible, very fast and allow for simulating multi-species and their ecological interactions. SFS_code is less flexible.

My only concerns about NEMO are that each individual dies after reproduction and that there can't be more than 256 alleles at the same time.

SFS_code limits the number of alleles to 4. Nemo limits the number of alleles to 256 (but you can also use phenotype and have a float number at each locus). SimBit uses either bi-allelic loci, a block in which the number of mutations are counted (max: 256 mutations; this is different than 256 alleles per say) or continuous float number at each locus for the phenotype. Do you really need more alleles? Why can't you just use several loci to represent a sequence with multiple alleles? Real biology can only have 4 alleles in its smallest locus! SimBit uses a bit-per-bit system. If you need, say 50,000 alleles, then you just need to ask for a sequence of 16 bits (2 bytes) and you can get $2^{16} = 65,536$ alleles.
If you need more alleles in a single block (which is a little surprising to me a priori), you can probably just ask the author and he might accept to helping you if you convince him it is of interest to add this feature (he might also be more willing to help in exchange of authorship if you need more support).
Hope that helps. Good luck!

Well, thank you a lot. You saved me a lot of time.

Though I haven't found a tool that fits the task well enough, the list provides several links to R and C packages that would save me a significant amount of effort needed to develop one myself.

I can't quite give you more advice as I don't exactly know what you're looking for. But you can simulate a whole bunch of things with NEMO and get the genes of every single indivdiuals afterward. NEMO is not super efficient for simulating an actual string of A,T,C,G (although it is feasible) but maybe something like SFS_code or SimCoal would do the trick. You might want to download Nemo (free of course) and run the example simulation that goes with the package you could get a better sense of what it does.

My only concerns about NEMO are that each individual dies after reproduction and that there can't be more than 256 alleles at the same time. Coalescence based techniques reconstruct genealogies, rather than inflate the sequence space. In my lab we are working on the structure of sequence spaces and we've found several hollow regions. Math-wise the problem is so complicated that we think it's better to try maximum-likelihood to estimate possible parameters of bacterial sequence evolution that could've led to this hollows. Hence the need in a simulator of forward sequence evolution.

@EliKorvigo I reread your comment and realized I might have not addressed it correctly. I edited my answer. The comment is now 2.5 years old. I am curious, what did you end up doing?

I'm terribly sorry I've totally missed your New Year's eve comment. I've ended up implementing a parallel basic evolution model with (almost) unlimited population size in C.


text:
What is this plant on the photos?
Found in Kustaankartano, Helsinki, Finland in June 2015.
(for closeup view, copy image URL and open in new window and then zoom in)

could we get some close up pictures of the flower? I'd suggest it may be some kind of clematis (is it a climbing plant?)

@rg255 I don't have them from another angle, unfortunately, however you can copy the image URL, open it in a new window and watch the photo much closer than SE shows.

@rg255 what about climbing, I've added one additional photo to the question, please check it out. It seems to be that it's climbing on a metallic netted fence.

It is one of the many species and hybrids of Clematis that is grown in gardens. It looks similar to the common Clematis alpina but might be a closely related species or à hybrid.

(Picture of C. alpina fr.o.m. Wikipedia)

Would have been my guess too. Those stems look pretty diagnostic. Do you find that quite often the answer to the question "what's that interesting blue-flowered vine?" is Clematis? :)

@Oreotrephes Hehe, yeah, it's often a good bet, at least in European gardens.

source: Stack Exchange

license: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/3.0/

sort: votes


text:
How should species density be calculated for a clumped distribution?
Lets imagine 5 plots of different size are sampled for a target species:
plot#  count  area(m^2)  plot_density
  1      1       5         0.2
  2      3       2         1.5
  3      0      10         0.0
  4      5       1         5.0
  5      2       6         0.33

What is this species' density?  I see two ways to calculate the density that give completely different values.
The first way averages the density at each plot:
$$
\frac{\Sigma(\frac{count_i}{area_i})}{5} = 1.41/m^2
$$
This seems OK but it doesn't do much to control for changing plot sizes.  For example if plot 3 above was 10 times larger the density would still be the same.  In a situation where plot sizes are determined by the environment (say, under natural cover objects) this seems less than ideal.
The second way totals the counts and the search area and divides them:
$$
\frac{\Sigma(count_i)}{\Sigma(area_i)}=0.46/m^2
$$
I prefer the second method because it seems to describe the animals' density more accurately. And I would be happy to use the second method, but my issue is that I am unsure how to calculate a summary statistic for method two since a mean was never calculated.  Method one gives a standard deviation of 2.1. What is the standard deviation for method two??
One possible solution I've come up with involves breaking the larger plots into 1m^2 plots and dividing the number of animals across those smaller plots.  So now I have 24 1m^2 plots with the following "counts":
plot#  count  area(m^2)  plot_density
1-5      0.2       1         0.2
6-7      1.5       1         1.5
8-17     0.0       1         0.0
18       5.0       1         5.0
19-24    0.33      1         0.33

Now, using the first equation above I get:
$$
\frac{\Sigma(\frac{count_i}{area_i})}{24} = 0.73/m^2
$$
With a standard deviation of 1.26.
Is this a reasonable approach?  Is there an established solution to this problem?

To me, there are two issues that are mixed up here (if I understand you correctly). First, do you want to estimate the mean and variance for a statistical population (i.e. to characterize a larger population by independent samples), or do you want to calculate the actual density for a particular area, where you have counted all occurences in that entire area (but maybe divided the area into subareas out of convenience when counting)? This is not clear from your question.
In the second case, your second option of pooling counts and areas is suitable. However, then you have only calculated the actual density in that particular area (ignoring issues of detectability of the organism when counting), and you cannot draw any inferences about a larger statistical population.
If your aim is to draw inferences for a larger statistical population, a start is to calculate the mean and standard deviation (sd) of your sample. In that case, I assume that your samples are chosen randomly and independently from a larger statistical population. Your first option is then the right approach. However, since your samples have different sizes, you might want to attach more weight to larger samples, since they can be assumed to better describe the population average than smaller samples. This is called a weighted mean.
Generally, the weighted arithmetic mean is defined as:
$$\bar{x}_w = \dfrac{\sum_{i=1}^nw_ix_i}{\sum_{i=1}^nw_i}$$
where $w_i$ are the weights for each sample ($x_i$) and $n$ are the number of samples. Using this formula, you will arrive at exactly the same value for the weighted mean as you calculated in your second attempt (0.458), if you use the areas as weights.
The weighted standard deviation is a bit more problematic, in the sense that there doesn't exist one single standard way of calculating this. However, a commonly used formula is:
$$ \sigma_{w_1} = \sqrt{ \frac{ \sum_{i=1}^n w_i (x_i - \bar{x}_w)^2 }{ \frac{(M-1)}{M} \sum_{i=1}^n w_i } },$$
where $M$ are the number of nonzero weights. Other definitions of the weighted standard deviation can be found at Wikipedia: weighted sample variance.
Another version is defined as (called "reliability weights" on the wiki page):
$$ \sigma_{w_2} = \sqrt{ \frac{ \sum_{i=1}^n w_i (x_i - \bar{x}_w)^2 }{  \sum_{i=1}^n w_i - \frac{\sum_{i=1}^n w_i^2}{\sum_{i=1}^n w_i}}},$$
If I haven't made a mistake, these will give the standard deviations 1.15 and 1.22, using your data.
As for the biological interpretation, all calculations of average densities indicate a clumped distribution, since the coefficient of variation (CV) is larger than 1 (CV = sd/mean), but more so if you use a weighted mean. The CV using the arithmetric mean is 1.5, while it is 2.5-2.65 for the weighted mean, which is reasonable since you are giving more weight to the large area sample with a zero count. However, I should also note that you should be cautious about using a weighted mean when you have a strongly clumped distribution, since your run the risk that e.g. a large sample plot land on an area with no occurences, which might bias your density estimate. Generally, when you have a clumped distribution, you need to sample more intensively to get a good estimate of average density, and many small samples is often better than a few large ones.

Great answer, thank you! If anyone else comes along and finds your answer helpful they might also be interested to learn the r package SDMTools has the functions wt.mean() wt.sd() and wt.var(). wt.sd() uses the more common equation that you described above.

After finding more time to consider your answer I have minor correction on your answer and a follow-up question.  The weighted mean for the above example is 1.16 not 0.458 like you say. And now consider these three plots: Count: 1, 1, 20; area(m2): 1,1,20.  I would say the density is 1animal/m2 and the three methods in my original post agree.  But the weighted mean is 18.2.  That doesn't seem right. What are your thoughts on this?

If you get a weighted mean of 1.16 you are calculating a mean based on counts, not densities, and the same goes your your new example. It also doesn't make much sense to calculate the average number of occurences (counts) for plots of different sizes, which is what you have done in your example in the comment. If you calculate a mean (weighted or otherwise) based on densities you will get 1animal/m2.

Yep you're exactly right.  Of course you would use the densities when calculating the weighted density. How I didn't see that before is beyond me.  Thanks again.

@AdamC You're welcome. Note that I've added a bit on the biological interpretation, and some issues when using the weighted mean for clumpted distributions.

all_licenses: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/3.0/


text:
Is the autonomic nervous system only activated by internal stimuli?
My professor claims that the autonomic nervous system is only activated by stimuli from organs but I really feel like I've read that it can be activated by outside stimuli, although I'm not sure what receptors that would be.
Is my professor right?

my bet is that you should examine specific pathway. Saying in general that ANS is excited only from within doesn't mean much, to my taste. ANS actions are mediated by CNS so one can say that environment does have influence on ANS

Short answer
External stimuli can drive autonomic responses.
Background
The autonomic nervous system is a visceral sensory and motor system. The viscera are the internal organs. Virtually all visceral reflexes are mediated by local circuits in the brain stem or spinal cord (Fig. 1). It is one of two major subdivisions of the nervous system; the other being the somatic (or voluntary) nervous system. The main distinction between the two is that the latter is involved in voluntary and conscious actions (e.g., movement, perception and cognition), while the autonomic nervous system is engaged in involuntary processes, such as regulating heart rate, breathing and bowel function (Fig. 1).

Fig. 1. Autonomic nervous system. Source: Austin Community College
Although by far most stimuli driving autonomous responses come from within (think muscle movements driving bowel movements, increased heart rates during physical exercise), outside stimuli do directly impinge on the autonomous system.
For example, the cilliary muscle in the iris is under the control of light coming into the retina. The pupillary light reflex test is based on this principle, and is routinely used to diagnose damage to the visual system. Also, production of saliva is controlled by external stimuli (smell and taste of food and see the striking example below from @anongoodnurse). And technically, the entire digestive tract can be considered to be outside the body, as it is continuous with the skin and lining of the mouth. Hence, the mechanical stimuli impinging on the intestines by a moving bolus of food that drives the waves of peristaltic movements can, arguably, be considered to be caused by external stimuli too. Lastly, the fight-flight-fright response involving the hypothalamic-pituitary-adrenal gland (HPA) axis is often initiated by external stimuli (loud sudden noises etc.).
The most important characteristic of the autonomic nervous system, as said, is that the autonomous responses are not voluntarily controlled. While you can decide to look or track a visual stimulus, you cannot decide to move the bolus of food back up to your stomach, or voluntarily constrict your pupil. The way the input is received may not be the best way to distinguish it from the voluntary nervous system.
Didactic answer
If you are left with questions, the first place to go to is your professor.
Reference
- Kandel et al. Principles of Neural Science 4th ed. (2000). Ch.49 The autonomic nervous system and hypothalamus. p.960-81

I wonder if the professor meant it on another level: that the autonomic nerve system does not have sensory neurons connected to the outside world. The examples you cite are ultimately caused by external stimuli, but as far as I remember, visual and olfactory perception is not part of the ANS. This is of course a nitpicking distinction and not meant to make your answer look bad (I liked and upvoted it), but for a student who is trying to find their way through complex material, easily delineated distinctions are good signposts, so it could be an explanation for why the prof used the statement.

@rumtscho - I have edited my answer to accommodate your comments. I agree that questioning your professor's words on SE may not be the best practice. In fact, my first short answer was always listen to your prof. I removed it, because lectures generate questions so it is logical to be left with questions. Further, there are always exceptions to the rule, that is fundamental to biological sciences. Please check my "didactic answer" and I hope that suffices your critique on a student's needs.

@AliceD - Hmm... good answer, but professors are often enough wrong that I think SE is a great place to double check. One fun trick (to which you alluded) is to cause saliva to flow by telling a patient to imagine biting into a lemon, because this has become a conditioned response in adults.  When there is a parotid duct obstruction, the parotid gland will swell before our very eyes., making the diagnosis obvious (and easy). Sensory nerves in the tongue for sour cause salivary production. There are other examples.

@anongoodnurse - Thanks for your comment and your interesting addition directly from medical experience. I made my answer stronger, and although leaving the didactic answer in place, I made it less binding. BTW: mod nominations at Bio are open :)

I saw that. You already have my vote!

@anongoodnurse - thank you :) But I was in fact hinting that you should nominate yourself!


text:
Pigmentation in Jaundice
My textbook says that the yellow appearance of a jaundiced person is due to the accumulation of bile pigments in the skin. I am unable to understand why these pigments are not deposited in a healthy person? Please help.

Because our liver breaks them down and makes sure they are excreted.

@Chris, my question was why couldn't the breakdown(that you are referring to) happen in a healthy person?

The breakdown does happen in a healthy person which is the reason they are not getting yellow.

The yellow color comes from the accumulation of Bilirubin in the body. Bilirubin is a breakdown product of hemoglobin (contained in erythrocytes, also known as red blood cells, and responsible for the oxygen transport in the body). (This is also the reason why bruises get colored, as the blood which was released into the skin gets removed.) As erythrocytes live only for around 120 days, the hemoglobin needs to be recycled at the end of the life cycle, as free hemoglobin is a pretty problematic substance.
The recycling of the erythrocytes happens in the liver, the iron is recycled, and the hemoglobin is split in the globin (which is further broken down into the amino acids) and the heme which is oxidized by the heme oxidase to biliverdin and further by the biliverdin reductase to bilirubin.
As bilirubin is not water soluble it is bound to albumin and released into the bloodstream for the transport to the liver. In the liver this process goes on further until the bilirubin is water soluble and can be released with the bile into the intestine where it is finally excreted. See the image (from here) for an illustration:

If your liver is not working properly (for example due to an infection) the break-down process of the albumin-bound bilirubin is reduced or doesn't work and the yellow bilirubin accumulates in the body until the skin gets yellow. So this indicates a liver malfunction and is usually nothing to take easy.


text:
Where is ATP used in Phosphorylation?
In the section describing the Calvin Cycle, under the subheading "Reduction",  my textbook states that:

The steps involve utilisation of 2 molecules of ATP for Phosphorylation...

I could not get where is ATP used in Phosphorylation (ATP synthesis by cells)? Please help me to understand.

This is a homework question. Even though you got your answer, please put in some effort on your part before asking a question. It is a good practice for everyone.

In biology phosphorylation marks the addition of inorganic phosphate groups to proteins or other organic molecules. The phospho-group usually comes from ATP which is converted into ADP in this process.
In the context of the Calvin Cycle there are two positions where molecules get phosphorylated. The first is the phosphorylation of 3-phosphogylcerate to 1,3-bisphosphoglyerate, the other is the phosphorylation of Ribulose 5-phosphate to Ribulose 1,5-bisphosphate.
See the image (from here):

Thanks for the help! so is it incorrect to say that phosphorylation is synthesis of ATP (since you said that the ATP is converted into ADP in this process)?

Yes, this is incorrect. Synthesis of ATP is usually referred to as "ATP synthesis".

@Abdullah, perhaps your confusion about the meaning of 'phosphorylation' in relation to ATP synthesis is because you may have previously learned of a process called 'oxidative phosphorylation' (sometimes abbreviated to OxPhos, or similar) that occurs downstream of the Kreb's Cycle--and is a mechanism cells use to synthesize ATP? Just a thought. The typical and predominant usage in lectures, textbooks, and papers is in Chris's answer that you accepted. The acid anhydride ATP donates its gamma phosphate to some other molecule in a reaction catalyzed by an enzyme.

Also substrate level phosphorylation…


text:
Can androgen-insensitive genetic-males get pregnant?
According to the linked report, people with androgen insensitivity syndrome appear as male, but have both feminine and masculine outer genitalia.
However, I was not able to find out whether their internal reproductive system is functional, i.e., if these people have a functional uterus.
Is it possible for such a person to get pregnant?

A "bug" is basically an insect or a computer software fault. Use proper terminologies. Please try to put in some basic effort. I could have simply said "no humans do not self-fertilize", but it is important to follow some basic discipline. Since you are a programmer and an active SO user, you would perhaps react the same way if I just ask a question without even attempting to code.

I specified the pathology in the question according to your linked report. Without mention of a pathology this question is too broad to my opinion. However, feel free to roll back if it doesn't reflect your original question.

Short answer
People with androgen insensitivity syndrome do not have a functional uterus and cannot bear a child.
Background
Androgen-insensitive genetic males may develop female genitalia and internal female reproductive organs. However, in both partial androgen insensitivity syndrome (PAIS) and complete androgen insensitivity syndrome (CAIS) the uterus is absent, or is present only in a rudimentary form.
The reason for the absence, or rudimentary form of the uterus is the fact that Mullerian inhibiting substance (MIS), produced by the Sertoli cells of the fetal testes is present. MIS controls the regression of the mullerian ducts, and in its absence in XX females, the mullerian system develops into the uterus, oviducts, and upper vagina. However, in XY individuals with CAIS (and likely also with PAIS), MIS is produced normally by the testes resulting in inhibition of the growth of the uterus and fallopian tubes (Oakes et al., 2008).
Reference
- Oakes et al., J Pediatr Adolesc Gynecol (2008); 21: 305-10

@mdperry - you cannot tell by looking who is an XY female, and your comment is misleading. I am not tall, and the two XY females I've known as friends are diminutive and certainly not fashion-model material. They are, in fact, no different from me in appearance, except they can't reproduce, and they are usually on hormone replacement therapy.


text:
Falconer & Mackay population mean calculation & genotypic values
I am reading chapter 7 of Falconer and Mackay where they give a formula to calculate the population mean as a deviation from the heterozygote genotypic value.
As an example, imagine a one locus two allele trait, with alleles $A_1$ and $A_2$, with respective frequencies $p$ and $q$, and genotypic values ($\theta$) of $A_1A_1 = \theta_{1,1} = 0.5$, $A_1A_2 = \theta_{1,2} = 0.4$, and $A_2A_2 = \theta_{2,2} = 0.3$.
The mean is given by
$ M = a_1(p-q) + 2pqd$
where $a_1$ is the deviation between the genotypic value of $A_1$ homozygotes and genotypic value heterozygotes, and, because $d=0$ (the dominance deviation), this equation reduces to
$ M = a_1(p-q)$
For my above example, if $p = 0.7$ then $M = 0.04$ and the population mean should be
$\frac{1}{2} \theta_{1,1} + \frac{1}{2} \theta_{2,2} + M = 0.44$
$\frac{1}{2} \theta_{1,2} + M = 0.44$
Is this correct?
What is the correct term and notation for the genotypic values (which I termed $\theta$)

You can't ping a user that hasn't participated on the post, they simply won't get the ping


text:
From where can I download the database created by Hsu et al for MORFs?
I am working on an independent research project for which I require access to the MORF database created by Hsu et al. From where can I download the database of MORFs of IDPs and their binding partners that was created by Hsu et al.?
I would appreciate a link to that page. I have searched a lot of keywords on Google but I could not come up with it. I might have, but I'm not too certain of it. I have already wasted 3 days searching for it.

In the future, it would be really helpful if you at least included a link to the paper by "Hsu et al." We can't read your mind. Secondly, the contact information for the corresponding author is included in the paper for a reason. If a careful reading of the paper and all referenced supplemental material doesn't answer your question, then contact the author directly. Most scientists I know are very happy to talk about their research and answer any non-trivial questions you may have.

I'm voting to close this question as off-topic because it is not about biology.

@MattDMo I got access to the database about a few hours after asking this question but I forgot to come and close this question. As for the question details, yes I completely agree with you. I made quite a few errors due to haste in typing the question. I'll make sure I do not do that again

I'm assuming you're looking for the dataset that was used to train MorfPred from this paper? http://www.ncbi.nlm.nih.gov/pubmed/22689782
There's a link to it on this page: http://biomine-ws.ece.ualberta.ca/MoRFpred/index.html
If you're referring to another paper, it is likely they used the same data set. The best way to find the supplementary data, is to read the paper thoroughly - this is a better approach than a Google search! Keep in mind though, that sometimes the data set is not publicly available and you must e-mail the researcher in question to request it.


text:
Additive genetic variance $V_A= 2pqa^2$ ... what is $a$?
Reading Falconer and Mackay looking at equation $V_A = 2pqa^2$ (equation 8.5, page 126).
I am trying to write a clear explanation of how additive genetic variance is defined and calculated, but I keep getting twisted up in the books I am working from.
Just what is the term $a$ and how is it calculated?

can you add a little context or provide a link to that article?

there's not much more context one can add, but I've linked to the book

Actually I could not access the book. Though you got an answer, I would suggest that you put up a snapshot of the page so that other users may find the question useful.

This information is given in p.118.
In essence $a$ represents the deviation from the population mean due additive genetic effects.
Say a trait is controlled solely by locus X with the effect alleles X impacting the phenotype Y in an additive manner (no dominance and epistasis effect), then we can calculate the effect size of carrying the X allele, called $a$, and representing the distance from the population phenotypic mean of homozygote XX. Based on this definition, the distance from the mean for individual with the following genotype is given by:

XX: $a$
Xx : 0
xx: $-a$

This makes more sense using the genetic model:
$P=G+E=BV+I+E=A+D+I+E$ (parameters listed at the bottom)
In the example, the locus has only an additive effect then D=0 and assuming there is no epistasis then I=0 as well which gives $P=BV+E=A+E$ therefore $BV=A$
In this case $a$ can be computed directly from the BV, with p=f(X) and q=f(x), as:

$BV(XX)=2qa$
$BV(Xx)=(q-p)a$
$BV(xx)=-2pa$

Now for the complete case of both additive and dominant effects, i.e. $BV=A+D$, you can refer to table 7.3 in p.118 which describe that XX individuals are given the value $a$, Xx -> $d$ and xx -> $-a$ with the corresponding relationship to the BV given by (care here it's not $a$ but alpha $\alpha$):

$BV(XX)=2q\alpha$
$BV(Xx)=(q-p)\alpha$
$BV(xx)=-2p\alpha$

with $\alpha=a+d(q-p)$

P=phenotype
BV=A+D=Breeding value
G=Genotypic effect
E=Environmental effect
A=Additive genetic effect quantified by $a$
D=Dominant genetic effect quantified by $d$
I=Epistasis effect

Thanks @cagliari that is a really good way of explaining it - are you studyin quantitative genetics? (I am in the chat room)

@rg255 Thanks! Yes and no, just finished a pop gen class where we used Falconer & Mackay intensively so this is still very fresh in my mind but my main research interest is actually in RNA/DNA structure prediction.

Cool the I'm stuck firmly in a popgen group - Hartl & Clark only, I had to do my own Q.G book clubs... I've realised where I was getting stuck too, it was $a$ becoming $\alpha$ that was throwing me off, but seeing equation 7.5 I see that $a = \alpha$ when $d = 0$.


text:
What bacteria shed harvestable energy from root zone organic matter?
I recently heard of a successful effort that harvests energy from soil at plants' bases due to apparently bacterial breakdown of wastes from the plants:

Via photosynthesis a plant produces organic matter. Part of this
  organic matter is used for plant-growth, but a large part can’t be
  used by the plant and is excreted into the soil via the roots. Around
  the roots naturally occurring micro-organisms break down the organic
  compounds to gain energy from. In this process, electrons are released
  as a waste product. By providing an electrode for the micro-organisms
  to donate their electrons to, the electrons can be harvested as
  electricity. Research has shown that plant-growth isn’t compromised by
  harvesting electricity, so plants keep on growing while electricity is
  concurrently produced.

Apparently there is more research here, and this was particularly interesting (a PDF) as it states more clearly that we are indeed talking about bacteria:

The PlantPower concept is based on the cooperation of plants and
  microorganisms to produce in-situ electricity. Plants take up carbon
  dioxide and water and capture light energy. This energy is stored in
  the chemical bonds of sugars produced, using carbon dioxide and water.
  Part of this chemically stored energy is transferred to the roots of
  the plants. This energy present in the root zone can then be captured
  by the so-called electro-chemical active bacteria. These organisms are
  capable to oxidize the organic matter present in the root zone and
  transfer the energy rich electrons to an electrode. The energy carried
  by the electrons can be used as electrical energy, after which the
  electrons react at another electrode with oxygen to form water.

I would like to understand this process as far as what bacteria or "micro organism(s)" are being leveraged here.  It seems that they are supplementing the soil with some kind of bacteria that acts in this specific way of releasing additional electrons that can then be harvested.
Any idea as to what kind of micro organisms or bacteria have this kind of reaction wherein they release or excrete harvestable energy (and apparently CO2 + H+) upon consumption of plant "excretions" (C6H12O6)?
After doing a bit more digging, I have found some, perhaps, interesting information from Wikipedia:

What ultimately confuses me here is that the other method noted earlier does not really emphasize any particular kind of plant and therefore I can't help but think that perhaps there are some other more useful bacteria for this scenario...

Geobacter is one bacteria that seems to get a lot of attention in the field of developing biological fuel-cells.

I am interested in this. I will lay down the realities of this plan that are probably implied; (the issues with conservation) Persuading large conservative bodies to switch (politics). Large scale farming and conversion of electric current. Nitrogenous waste conversion, It could replace industries that make ammonia therefore Haber process gets replaced. Grass plants are more reliable but Colonization of invasive virus species destroying crops is the reason mono culture went towards DDT and pesticides, besides the fact DDT is a war invention. It is interesting also if mushrooms could supply the

Short answer
It is known that a great variety of soil micro-organisms is able to use solid anodes as electron acceptor for their metabolism, thus creating some electrical current. Geobacter spp., Clostridia and Deltaproteobacteria are among them (1). However, I could not find any extensive list of these microbes. It is likely that a large number of these microbes is not identified yet.
A review on Plant Microbial Fuel Cells was recently published and might interest you (2).
Long answer
Plant Microbial Fuel Cells are a special case of Microbial Fuel Cells
The system presented at your first figure is being developed by the Dutch company named Plant-e and falls within the concept of Plant Microbial Fuel Cells (PMFC).
Simply put, a Microbial Fuel Cell (MFC) is a device in which chemical energy contained in chemical bounds of organic substrates is converted to electrical energy by microbes. Microbes consume the organic substrates while using the anode as the final electron acceptor of their metabolism. Microbes capable of using anodes as electron sinks for their metabolism are called electrogens.
PMFCs are a special case of MFCs, taking advantage of the microbe/plant interactions. Organic substrate is supplied to microbes directly by the plant and you get two benefits: plant biomass and electricity production.
Complex interactions between plant and microbial communities
As you noted in your last sentence, no particular species of plant was cited. Indeed various PMFCs strategies were implemented, based on various species of plants, such as Glyceria maxima, or Oryza sativa (rice) (2). The plant species is not the only key factor for efficient PMFCs: the nature of the organic substrates provided by plants to microbes, the soil's characteristics, the electrodes' characteristics, and the nature of the inoculum are crucial factors too.
Regarding the microbiological aspect, it appears that pure culture (single species) PMFCs are not necessarily the best alternative. Plants provide microbes with a variety of chemical compounds, which may not be all consumed by a single species. Only a well adapted microbial community is likely to be able to take advantage of all these compounds and to include efficient electrogens. The path from plant products to electrical current is more likely to be a complex microbial food web, rather than a single species' metabolism. In practice, most studies seem to rely as soil as the incolum (2).
So, concerning your initial question: we might not have identified yet every soil electrogens. Several are, however, already well known, such as Geobacter sulfurreducens. But, what is important is that most (P)MFCs strategies rely on microbial communities as a whole, in a 'black box' way.

References
(1) Dunaj, S.J. et al. Relationships between Soil Organic Matter, Nutrients, Bacterial Community Structure, And the Performance of Microbial Fuel Cells, Environment Science and Technology, 2012, 46 (3), pp 1914-1922
(2) Nitisovarut, R., Regmi, R., Plant microbial fuel cells: A promising biosystems engineering, Renewable and Sustainable Energy Review, 2017, 76, pp 81-89


text:
Following DNA replication during S-phase of the cell-cycle, are all genomic regions subjected to the same stringent level of DNA-Repair?
To my (limited) understanding, there are 2 main ways that mutations can occur in DNA: Environmental (UV, etc) and mistakes during cell division.
I was wondering if there is a mechanism that can give priority to certain genes to be accurately duplicated. Some sort of trigger that says "double-check this specific gene before continuing with the duplication".
And if there is such a mechanism, then I wonder if there is some sort of dependency system for genes that control groups of other genes. So that if a certain gene "activates" the double-check trigger, it would automatically add that trigger to the group of genes which are affected by it.
Thanks.

All DNA is proofread during replication. It is also constantly monitored for damage.

are all regions have same mutation rate? bonus points: independent of sequence (only location). E.i., are there markers of super-stable sequences

Can you add some details. Your exact question remains unclear to me. Gene duplication is not essentially a regulated process.

No such mechanisms as far as I know. Some DNA regions are harder to repair, notably repeated sequences but I am not aware of any mechanism giving a specific higher priority for repair to any genes. Also I think you want to use the term "replicated" and not "duplicated". @aandreev while interesting, mutation rates seem irrelevant for this question.

So far, the known mechanism of DNA repair is to recognize mismatches or damaged nucleotides by enzymes surrounding DNA rather than by scanning along DNA. Therefore double check could not happen under such conditions.

DNA mutations arise from either replication errors or the effect of mutagens. Cell division can go into what is known as the G0 phase where division is put on hold. This state is also referred to as a quiescent state (see the related genes here), which is distinct from senescent states where damaged cells are prevented from replicating by, for example, the p53 gene family. The proteins of these last mentioned genes are up-regulated to suppress uncontrolled division in cancerous cells, while genes controlling the G0 phase act under normal physiological conditions, unless these related genes are deficient themselves. Carcinogens are a subset of mutagens that typically affect genes involved in the cell cycle.

I was wondering if there is a mechanism that can give priority to
  certain genes to be accurately duplicated. Some sort of trigger that
  says "double-check this specific gene before continuing with the
  duplication".

A mechanism that would say: "double check this part, it's important" isn't known to me neither, but if I understood your question correctly I think the answer to your question lies somewhere else.
Let's start with the mutation rate you mentioned, some sequences are harder to replicate by DNA polymerase (it makes more mistakes there, such as in highly repetitive sequences, methylated cytosines etc.), so the mutation rate is higher in these segments (they are called mutation hot spots), but otherwise the DNA polymerase makes errors randomly, throughout the whole genome. These are in better case repaired. No special care given, repair polymerase tries to repair everything.
Now, to my point. What if the repair fails? In genes, that are critical to cell survival, development of organisms etc. (for it's life) you really see less mutations (if any) present. (and I believe this was the reason for the question, correct me if I am wrong.)
Moreover, these places in genome can be highly conserved between different organisms. More here. What this says is that nature doesn't allow cells/evolution/whoever to experiment (or make errors) on these positions. If mutation occurs here and isn't repaired, the cell simply dies and therefore you cannot see the mutation. The reason for death may be accumulation of toxic protein, which cell is not able to get rid of with disrupted enzyme, or lack of energy, because it cannot oxidize fuel... there are plenty possible lethal outcomes. For some genes, the cell can substitute their product by another protein, but some are essential.
(In multicellular organism can substitute for it, unicellular won't care because it's dead already)
I believe this is the mechanism, by which cell can distinguish between important and "less important" genes you were asking for.


text:
One Sample, Different Methylation Values
I'm hoping that somebody here can explain this unusual result that I'm getting with some pyrosequencing data. I have bisulphite converted a few samples a couple of times over the years for different projects. Recently I was collecting some of this disparate data together and I noticed that when the same pyrosequencing assay was used on these samples, those that were converted later had decreased methylation at the CpG sites of interest compared to the initial bisulphite converted samples. The original conversion was in 2010 and the repeat conversions were in 2014/2015. The assays are the same and it's difficult to blame this on PCR amplifying one clone over another since I've done technical replicates on each aliquot. Even the 2010 converted sample has the same result in 2015 that it did in 2010 but this is different from the 2015 converted sample.
We are using the same kit (Qiagen) for bisulphite conversion and haven't made a major change to our procedure (we follow the Qiagen protocol without modification). Has anybody noticed this phenomenon before? Is it possible that stored DNA would experience methylation changes? It seems more likely that the issue is technical but I'm having trouble figuring out exactly what that technical issue is.

In experimental troubleshooting questions like these it is difficult to provide an exact answer unless you specify the protocol. You should also add details such as storage conditions, how old was the bisulphite solution etc, level of difference between the two replicates etc.

Thanks - I added some detail. I unfortunately can't say too much about our pyrosequencing assays until we've published.

So just to clarify: You have a sample, converted some of it, stored some of it. Sequenced some of the converted sample(A). Wait five years. Sequence some more converted sample(B), convert some of the stored sample, sequence that (C). So A and B match but C is less?

I looked into this and if the effect is real, which it sounds like it is, it may be 5-formylcytosine.
Methylcytosine oxidizes into hydroxymethylcytosine, which reacts with bisulphite exactly like you expect (identically). Hydroxymethylcytosine further oxidizes to 5-formylcytosine, which behaves like unmodified cytosine (i,e. deaminates to form U). TET enzymes mediate this behavior in vivo but I can't find any data on spontaneous methyl group oxidation or what the kinetics might be like. This paper relies on the properties of 5-formylcytosine outlined above.
You could use this paper to test if that's true. Reduction with sodium borohydride converts 5-formylcytosine back to hydroxymethylcytosine, which reads as methylated under bisulfite sequencing.


text:
What limits chromosomal length?
What are the upper and lower limits for chromosome length? Are these limits different in different species or kingdoms? If there is any limit, which cellular or molecular factors are reasons?

There is both an upper and lower limit which are species specific. The upper limit is caused by incomplete segregation of sister chromatids and subsequent trimming of the long arm(s). The reason for the lower limit is, as far as I can tell, unknown.

Upper Limit
Schubert I, Oud JL. 1997. There is an Upper Limit of Chromosome Size for Normal Development of an Organism. Cell 88:515-520.
Hudakova S, Künzel G, Endo TR, Schubert I. 2002. Barley chromosome arms longer than half of the spindle axis interfere with nuclear divisions. Cytogenet Genome Res 98:101-107.
Rens W, Torosantucci L, Degrassi F, Ferguson-Smith MA. 2006. Incomplete sister chromatid separation of long chromosome arms. Chromosoma 115(6):481-490.
In these papers they found that in mono- and dicotic plants and one animal, the chromosome arm length is limited to half of the average length of the spindle arm axis (the length between the centromeres). Due to the mechanism and its presence in both plants and animals, it might be reasonable to expect that other domains of life are subject to similar constraints. Schubert and Oud (1997) speculate that the spindle axis length is fixed species specifically.

These are confocal microscope images of dividing Vicia faba (bean) cells (Schubert and Oud, 1997). (a) Wild-type cell at anaphase. (b-d) When the chromosome arms are longer, the chromatids cannot be completely separated at anaphase. (e) Wild-type cell at telophase. (f, g) Longer chromatids cannot be separated sufficiently and a bridge forms between the two daughter nuclei. (h-j) The bridge is broken when the cell plate forms. (k) Wild-type cell at interphase. (l) Breaking of the bridge between cells with lengthened chromosomes creates irregularly shaped nuclei and micronuclei - these micronuclei represent loss of DNA from the genome and are eventually lost outright through further cell divisions.

Lower Limit
There is a lower limit, but it doesn't seem that anyone knows why, which makes it impossible to accurately define.

Schubert and Oud (1997): Data from various eukaryotic kingdoms indicate the existence of a lower limit of chromosome size for stable mitotic and meiotic segregation. Yeast artificial chromosomes do not exhibit normal meiotic stability until they are of a size between 120 to 500 kb which comes close to the order of size of native yeast chromosomes, in addition to functional telomere and centromere sequences (Niwa et al., 1989). For a Drosophila minichromosome, an island of complex DNA of 220 kb was shown to be necessary for proper centromere function (Murphy and Karpen, 1995). However, for stable mitotic and meiotic segregation an additional 200 kb of a flanking satellite sequence was required. For stable achiasmate disjunction of minichromosomes during meiosis in female Drosophila, 1000 kb of overlapping centric heterochromatin sequences are necessary (Karpen et al., 1996). Schriever-Schwemmer and Adler (1993) have shown that a minichromosome representing the small centric product of a translocation between chromosomes 7 and 15 of the mouse and comprising about 1% of the haploid genome got lost during meiosis in the female carrier organism while the corresponding product of a translocation involving chromosomes 4 and 8 did not. The latter was somewhat larger. Either this different behavior was due to the presence of essential genetic information on the small translocation product 4⁸, or the minichromosome 7¹⁵ was just too small for stable segregation, although it contained a complete centromere as well as telomeres. Finally, we have observed that the frequency of meiotic missegregation, but not of transmission to the next generations, is inversely correlated with the size of translocation chromosomes in Vicia faba (Schubert et al., 1986). Thus, the existence of a proper length range for eukaryotic chromosomes is obvious.
The physical reasons for the lower limit that still enables stable mitotic and meitotic segregation are less clear at present than the constraints for an upper limit of chromosome length

Of course, "present" in that paper refers to the late 90's but, in my admittedly brief search, I have been unable to find too much in the way of more recent results. The following paper seems somewhat promising, but I don't have access to it and can only quote from the abstract. It's not much more recent, either.
Schubert I. 2001. Alteration of chromosome numbers by generation of minichromosomes -- is there a lower limit of chromosome size for stable segregation? Cytogenet Cell Genet 93:175-181.

[A] lower size limit for a stable chromosome transmission … might be based, for instance, on insufficient lateral support of centromeres or on insufficient bivalent stability due to the incapability of chiasma formation.

I must say the effect of a chromosome being so physically long that the sister chromatids cannot divide anymore between the daughter cells is amazingly interesting. I wonder if there is any kind of quantifiable equilibrium between a genome length and the number of chromosomes, i.e. having "short enough" chromosomes yet not too many as the more of them the higher the chances of chromosomal nondisjunction events. (disclaimer, I couldn't find a link between chromosome number and occurence of disjunction events, was just a wild guess). Anyways thanks for this great answer!


text:
Why do some protocols require prewarming a liquid medium before inoculating?
For example, in this protocol for E. coli competent cell preparation, it says:

Plate 10 uL E. coli BL21(DE3) cells on a LB-agar plate; incubate overnight (12 hours).
Prepare 500 mL SOB medium so it can be pre-warmed in the 37 degrees Celsius incubator the day before.
Pick a single selected colony and inoculate in 5 mL of SOB medium and grow for 16 hours at 37 degrees Celsius and 250 rpm in a shaker.
Inoculate 500 mL of prewarmed SOB medium with 5 mL of the fresh overnight culture.

and then in the protocol for overexpressing proteins after transformation of these cells, I basically follow similar steps as above but this time, without prewarming the large-scale inoculation lqiuid medium. Why is that?

Dramatically changing the temperature quickly can shock cells, both prokaryotic and eukaryotic. This could, for example, cause quickly-growing log phase cells to arrest for a period of time, depending on the temperature of the large volume of media (i.e., RT or 4C).

Growth speed also changes by incubation temp. If you put large, medium, and small sizes of culture media kept at room temp into 37 degrees incubator, times when you get the desired density of E coli may vary among the sizes

It's because E.coli BL21(DE3) are competent cells. The competent is the key here as the cells were chemically treated so the transfection can be performed by heat-shock with high efficiency.
This means these bacteria are quite fragile, due to the chemical treatment, and therefore are very sensitive to both mechanical and thermal shocks.
Pre-heating the media simply avoids damaging the cells and maximizes the growth rate.


text:
Choosing a book to gain general knowledge about biology
I will be first year undergraduate at Physics department next year and last year I was at the Medicine faculty. I want do double major in the second year of faculty with Molecular Biology and Genetics. The intersection of biology and physics is my interest. I have two books about biology, first one is What is Life?-Mind and Matter by Erwin Schrödinger and the second one is This is Biology by Ernst Mayr. Which one should I read to acquire general knowledge about Biology?

The books you have
The books you cited won't help you to get a solid and general basic knowledge in biology. I haven't read those books but I think that This is Biology is a book of philosophy of biology written by a biologist and philosopher. What is life is a very influential book written by a physicist who more or less predicted the structure of DNA that was later discovered by Watson and Crick. You won't learn much of biology through these books.
Book Suggestions
This post offers book suggestions for general introductory biology. Here is another post that may also interest you
If you are interested in a specific field of biology, then you may want to ask for book suggestions for this particular field. I think that most biologists that come from a background in physics are in the fields of theoretical evolutionary biology, bioinformatics and biophysics (incl. molecular biophysics).
Here are a bunch of other posts asking for book recommendations that I imagine you may like reading about.

What physics knowledge can be applied to biology of organisms and ecosystems?
Book recommendations for algorithms used in evolutionary biology
Book recommendations for evolutionary models
How to learn biomathematics?
Book recommendation on population/evolutionary genetics?
Introductory literature for synthetic / systems biology?

And eventually you may want to read Value of mathematical models in biology although the post has been closed.
Of course, you might want to read a bunch of wikipedia pages such as Biophysics, Molecular Biophysics, Mathematical and Theoretical Biology, Biostatistics, Computational Biology, Bioinformatics.


text:
Neomycin resistance gene in E. coli stb3 cells
We use a lentivirus packaging vector called pLenti_puro_DEST. We use puromycin to select for the cloned gene in 293T cells. However, the pLenti_Neo_Dest has been discontinued, so I'm cloning a Neomycin resistance gene into the pLenti_Puro_Dest thus replacing Puromycin with Neomycin. I usually just grow up this gene in Stb3 or Stb4 cells from invitrogen. I'm wondering if I can select for cloned pDEST vectors in these cells using Neomycin or G418. According to the G418 manual, you can theoretically use it for bacterial selection, but I think that may be for libraries and not actually growing up the plasmid.
So is it possible to select for my plasmid in these cells by adding a little G418 to the ampicillin plate? Is this common? I can't find anyone selecting bacteria by Neomycin anywhere.
EDIT - The main reason I ask, is that the resistance gene is under a PGK promoter that I have no idea if it works in bacterial cells.

Mammalian promoters don't work in bacteria basically.

PGK is mammalian (murine) phosphoglycerate kinase promoter. It will therefore not work in bacterial systems. Why don't you select using ampicillin only? You can increase the ampicillin concentration if you want.
Regarding:

I can't find anyone selecting bacteria by Neomycin anywhere.

Kanamycin is generally used for bacterial selection. The Neomycin resistance gene works on Neomycin, Kanamycin and G418 (a gentamycin variant). The gene basically codes for an aminoglycoside phosphotransferase and different variants of the gene have different affinities for different aminoglycoside substrates. For a basic reference see this.

I do select with ampicillin, but any residual parent vector (from the cloning) would also contain ampicillin.

@jwillis0720 For that you have to do screening. You can do colony-PCR screening which is relatively faster than other methods. Also have self ligation controls to have an idea of how many colonies you need to screen. This is sticky end cloning right?

Plasmid  construction is a technique to get plasmids that carry your inserts from empty vectors carrying the same resistant markers. If you want to select with kanamycin/neomycin, you could take a bacterial expression unit of a kanamycin/neomycin gene. In fact, there are plasmids which carry kanamycin with both bacterial and mammalian promoters,

I will just screen by PCR. But it's gibson assembly.

@jwillis0720 not an issue. Just use the end primers and screen for the appropriately sized amplicon.

I understand, but the amplicon is so freaking big, it approaches the template vector size supercoiled.

@jwillis0720 Ohh okay..in that case RD would be better.. Or a colony PCR on a section of insert followed by RD

In this paper (Li and Elledge, Nature Genetics, 2005) it is shown that the PGK promoter works in E.coli but it has very little activity. So, I don't think it is suited to properly express resistance enzyme.
About Neomycin, many aminoglycosides resistance enzymes (not all) that work on Kanamycin works well also with Neomycin. I have personally selected E.coli expressing Kan resistance (from a pET vector) using Neomycin without problems.


text:
Gene silencing: relative or absolute number of methylated CpG sites?
CpG content varies a lot across the human genome. What does determine a promoter to be silenced through DNA methylation: the fraction of CpG sites that are methylated or the total number of methylated CpG sites? Any reference?

Total methylated sites in the genome or in the promoter?

In the promoter

Generally, robust silencing by DNA methylation requires that all or most CG sites be methylated and that there are a good number of CG sites to methylate.  (A 'good number' is relative to each species depending on CG content of the respective genome.)
A promoter with ~10% of its CG sites methylated would not likely be silenced.  However, the caveat with this is that one well-placed methylated CG can cause silencing or at least decreased promoter efficiency.  Also, many promoters are depleted for CG, which means they can be fully methylated, but still not have much absolute methylation.  These promoters can be active despite apparent 'hypermethylation' in the CG context.  Also, a caveat to this is that promoters with zero methylation can be silenced, even if they have CG sites.  Basically, if CG methylation is important in the regulation of a gene, it will have CG sites in functionally-important parts of the promoter.  The promoter will be well-methylated when the gene is silenced, but because placement of de novo CG methylation is not absolutely perfect, there is a high likelihood that there will be more methylation than is needed for silencing to be achieved.  A sort of overkill phenotype.
This paper from the Schubeler lab is a nice foray into the mammalian promoters, which they break into 3 different groups depending on CpG content, and then further analyze each group for CG methylation profiles.


text:
What determines the number of chromosomes an organism carries?
This is an extension of this question about What limits chromosomal length?.
I am wondering what could be the specific reasons behind the number of chromosomes an organism carries. In other words, what drives the number of chromosomes and is there anything "stabilizing" the number of chromosomes.
Of course I am not talking about multiple copies of the same chromosome, i.e. trisomy, but rather the reasons behind the partition of the genetic information into a defined number of chromosomes.
From the question I referred to, chromosomes have an upper length limit which is based on their physical size, i.e basically they must be shorter than half of the spindle arm axis and they probably also have a lower limit. Yet it is unclear to me if any non-random equilibrium exists between chromosome number and length other than just being bound by an upper and lower limit in size.
Also let's restrict this question to diploid organisms to avoid organisms such as the Oxytricha trifallax which has 16,000 chromosomes but is also ampliploid or plants such as wheat which can be tetraploid.
I tried to search in the literature and couldn't find a satisfying answer. Also I had in mind the example of the Hela cells which have completely shuffled chromosomes yet still exactly 46.

With more chromosomes, the cell would need more number of centromeres and telomeres (this unnecessarily increases the size of the genome). Also the metaphase checkpoint has to be more stringent to prevent aneuploidy. This is just a guess though. In general there would be a higher metabolic cost in maintaining a very large number of small chromosomes compared to fewer but longer chromosomes. Additionally meiotic crossover would be relatively difficult with smaller chromosomes

Practically, there is an obvious barrier to changes in chromosome number: haploid cells produced by diploids must contain exactly one copy of each chromosome (or chromosome section, or locus). Otherwise over- or under- expression of genes can lead to dosage problems. Across all of diploids some large groups show strong conservation of number of chromosomes but others seem fairly free to change the number.
An example of conservation are the Drosophilids (fruit flies) where for approximately 100 million years they have maintained five or six chromosomes (search for Muller elements). Incidentally, with a huge amount of intra-chromosomal shuffling of genes "Drosophila 12 Genomes Consortium", 2007, Nature.
An extreme example of changes in chromosome number can be seen between similar species of Muntjac deer . Where in a relatively short time period (of the order of a million years or less (?) - my guess), the number of chromosomes has reduced from 46 to six (Wurster, D. H., Benirschke, K., Jun. 1970). Someone may correct me but I think amongst vertebrates, mammals show particularly fast evolution of chromosome number relative to, say, birds (though more birds also have micro-chromosomes...).
The above changes (or lack of change in Drosophilids) appear to be independent of genome size - so that was a long way round to saying "It's not genome size".
It could be repetitive DNA. The centrosomes (where mitotic spindles attach) of mammals are full of repetitive DNA and for this reason have been hard to sequence accurately (Graphodatsky, A. S., Trifonov, V. A., Stanyon, R., 2011. ). These repeats evolve (in sequence and copy number) and may promote more or fewer spindle attachments. Drosophilids (in general) have fewer repeats but I don't know if this is a diploid-wide pattern [Research anyone?].

Very interesting, thanks! Do you have, by any chances, some direct references I can look at? What I would like to know is if it is solely random recombination that drove those changes which would indicate that the number of chromosomes in an organism would change randomly within some size constrains.

I've added a few refs that may serve as starting points.  When you say "recombination", do you mean recombination (the normal pairing and crossover of homoologous chromosomes during meisois) or chromosome-rearrangement (mutation leading to novel chromosomal structure)?


text:
Can photosyntetic cells be genetically modified to grow at a faster rate?
I hope the question is not silly, but I wonder if a sort of genetically modified plant "cancer" could be used for carbon sequestration. For example, cancer cells like HeLa have been reproduced in tons, and as far as I know their growth rate is very high.

You might be interested in reading about artificial photosynthesis.

Yes! Thanks, I was just reading that. I still think the question is relevant as we are relatively advanced in OGM, and tumors and plants already are there. It seems to me (I'm completely ignorant in biology) that it would be easier than reproduce cheap photosyntesis (i.e. cheaper than planting trees).


text:
Identify the plant
Can anyone tell me which plant or flower is this. I have found it in Darjeeling,  West Bengal, India.

Those are known as Rhododendrons, and I'd ID that one as R. hodgsonii.


text:
Do we need to drink less water in humid weather than in dry weather?
From my understanding, humid weather means that the air has a lot of water in it. Therefore, someone who lives in this kind of weather would be hydrating himself just by breathing.
Does that mean that, to stay hydrated, one needs to drink less water in humid weather than in dry weather?
Of couse, I know that humid weathers are usually hotter, which accelerates dehydration. But in this case, let's assume the temperature is independent of the humidity of the air.

I think it depends.  At least from personal experience, I seem to need more water in dry cold than humid.  And in hot, humid weather, I seem to lose more water by sweating simply because the humidity makes the cooling process less efficient than it would be in dry air at the same temperature.

@jamesqf You seem to lose more water in moist weather but actually you lose more in dry weather.

You don't get hydrated by breathing moist air. That may at best hydrate your respiratory tract. You should drink more water in dry weather because the body gets dehydrated more.
Humid weather is not hotter, it just feels more uncomfortable. But dry heat is actually more harmful, because it dehydrates you (not in the sense of extreme heat).

It all sounds plausible, but do you have references to back this up? Particularly, when I am hiking in humid Florida or Maryland I drink many liters of water a day. The thing is that sweat doesn't evaporate and it seems the body is simply only heating up. In dry heat, however, sweating is way more efficient as evaporative cooling is always more efficient in dry climates, such as in most parts of Australia. When I hike in Australia, I do not particularly drink more, on the contrary even, in similar temperatures. Anecdotal stuff, but still something to contemplate about, perhaps.

@AliceD I didn't see any reference. I'll look up; I am not sure if there is actually some review on this. At very high temperatures moist heat would be more damaging (because of specific heat of water) but drinking water per se just hydrates your body. I don't think it has a direct role in bringing down temperature unless of course you are drinking cold water.  This is all based on the physics of it. I'll look for a reference.

Not referenced, but MIT indicates it is unclear which climate makes you sweat more, http://engineering.mit.edu/ask/why-do-we-sweat-more-high-humidity

@AliceD You do apparently sweat more in humid climate because the water doesn't get evaporated. Dehydration is more in dry clmate.

Yes exactly my point: perspiring equals dehydration. The question then becomes, due to the inefficiency of perspiring in humid envrionments whether you perspire more in humidity, or whether it only seems so because it won't evaporate.

@AliceD You apparently sweat more because the sweat doesn't evaporate. The principle is similar to osmosis if you see. You may notice that though humid heat is uncomfortable, there are less incidences of heat strokes and heat cramps (anecdotal).

@WYSIWYG If I get you right, you're saying that we perspire the same in dry and humid weather. If that's so, then why does dry heat dehydrate us more than humid heat? Shouldn't they dehydrate us the same?

@Zenadix dehydration is not just because of perspiration. You would get dehydrated if you are in a dry atmosphere. Heat increases the evaporation rate. After a particular limit of dehydration perspiration reduces (see this article) but dehydration nonetheless happens. Imagine keeping a tomato in a dry condition versus keeping in moist condition. Tomato doesn't perspire but will nonetheless be dehydrated in the dry condition. I'll add this point to the answer in a while.

First of all, we sweat as a method of normalizing/reducing our body temperature caused by the situ temperature, outside of our bodies. This mechanism, is primarily completed with thanks to the Eccrine sweat glands - those that cover the greatest majority of the body and hence, are the most essential in this context.
Sweating is classed as 'evaporative heat loss' so, the correct conditions have to apply in order for the mechanism to effectively occur.
Humidity: The common perception is that we sweat less in hot and humid conditions since, "the sweat just sits there", inferring that we are perspiring at a slower rate. However, it's the complete opposite. This is because, in the stated environmental conditions, the ambient water vapor pressure is greater and resultantly, the body needs to increase the area effected by perspiration -- the sweat rate increases -- thus to match the same effect that you'd experience in hot, dry conditions. The affinity for the evaporation product to attach to molecules in the air is less due, to there being more water molecules already present in the air. It's a bit like how water in cells moves from areas of high potential to low water potential.
Dry:
The body can more efficiently cool itself in this environment. This is due to fact that dry air has a greater affinity for water molecules so, the sweat will evaporate quicker. Additionally, this has a knock-on affect on salivary glands and furtherly, the humidity of your essential respiratory organ, the lungs - causing dryness. This is very prevalent when breathing heavily e.g. in exercise.
This is very similar to hygroscopy.
To conclude, you need to drink more water in hot, humid environment than the contrary.
If my answer is too "generalized"; i apologize.
Citations:

Eccrine sweating
Jay Hoffman in Physiological Aspects of Sport Training and Performance


text:
Expected reproduction rate of a dandelion and/or fern
I am working a basic mathematical/computer simulation of an ecosystem, and I need to know the birth rate of each organism in my simulation. However, I could not find any data for how often a small, asexually-reproducing plant, like a fern or dandelion, reproduces. Does anybody know either a) the time to reach sexual maturity and the expected offspring of one dandelion or fern OR b) when starting with one plant, the expected living dandelion or fern plants after a year?
In other words, if I plant one dandelion or fern in a closed system (providing it with requisite water, food, sunlight, etc), how many can I expect to find at the end of the year?
These values don't have to be exact, I can adjust them later, but I need something to use as a baseline.

Dandelions may have "from 54 to 172 seeds per head and a single plant can produce more than 2000 seeds" (https://fyi.uwex.edu/weedsci/2002/11/12/dandelion/), but the number of inflorescences depends, for example, on the age of the plant. It should also be remembered that some individuals repeat the blooming in autumn, although it is not so abundant (usually only one inflorescence). That, what is called a dandelion (Taraxacum officinale) is a collective species, which actually consists of several hundred microspecies, which may differ in the amount of seeds produced. Thus, the given data is only quoted.

all_licenses: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/4.0/


text:
Leaves growing on a leaf of a Begonia
I've a Begonia species. Somehow it started to grow new leaves on top and right in the center of an existing leaf which you can see in the center of the image. Why is it doing this? Apparently it is not a disease because the Begonia itself is growing fast. Will the new leaves eventually become a new Begonia which can survive on its own?

The phenomenon you are observing is probably Adventitious buds. The plantlets can develop on intact leaves or on injured one.
Back to Begonia, this phenomenon usually occurs on injured leaves, so, probably, the plant was injured during movement/by insect or by other means and because of this started to create plantlets.

Will these plantlets eventually be able to survive on their own?

@wie5Ooma I think yes, definitely...

 African Violet plantlet growing from another african violet leaf that I'm growing. This one is growing at the stem and leaf junction. I also have another leaf that has three plants starting to grow along the bottom edge of a leaf. If I remember from my high school days in horticulture class,(when I was in school with Moses), this probably occured when I cut and potted the leaf and scratched or rubbed the leaf thereby stimulating the cells and causing growth. The article before mine gives you the scientific version of this phenomenon.

Hi James. Welcome to [biology.se]. I want to point out that this post does not really answer the question; it is more like "I have this problem too". If at all it does answer the question to some extent, it is based on anecdotal evidences. Can you please cite a relevant source (perhaps a horticulture manual, book or a research article) to support your claim?


text:
Will the heart keep beating if it's separated from the body?
The heart is a vital organ in our body, as it drives blood circulation. I was wondering if a heart keeps beating if it is separated from the body? If yes, then why?

This is more like a homework question. Please put in some effort towards answer.

@WYSIWYG I seriously have no idea if it is homework question or not but before asking I do searched in google which states that heart has its own electrical impulse but I wanted to know that is there anything else happen while our heart is seperated. I have never face this situation but for sure you are much experienced than me so I'll make sure to show my efforts when I will ask any sort of homework question next time . thanks

You have heard of heart transplants I suppose. What I am saying is that you should try avoiding one line questions. Add whatever you got from google and make the question precise. You can check some meta posts on "homework". Homework is not in strict sense something that you get in your school. It is in the sense of asking - "have you done your part of work i.e. homework, before asking others".

That's true no doubt on that .. I'll make sure to make question precise.

Short version
The heart  has the ability to beat independently of the brain as long as it has oxygen. The heart will eventually stop beating as all bodily systems begin to stop working shortly after brain death. Remember the heart can beat, but your diaphragm and lungs won't. Hence the cardiac muscles undergo asphyxiation and die off. However, immediately after death, there is enough oxygenated blood in the body to keep the heart pumping for a while.
Long version
There is a simple explanation why.
As you know every muscle in the body has to receive stimuli from the neuromuscular junctions (and subsequently the nervous system) in order to contract.
The heart is a bit different in that it is not regulated by the brain, but the regulatory mechanisms lie within the heart itself. The heart conductive system contains a special group of cells called the pace maker cells (SA node) that fire at regular intervals and cause the heart to beat.

Each heart beat is triggered by an electrical pacemaker - a group of cells in the heart that have the ability to generate electrical activity. They cause electrical impulses to spread over the heart and make it contract.
The largest natural pacemaker of the heart is called the sinoatrial or SA node and is found in the right atrium. From it, specialised groups of cells that carry the electrical charge lead off to the rest of the heart.

(Taken from here)
The brain regulates the rate of the heart beat sure, but it does not send the signals that cause the heart to beat in itself.
In short, even after disconnection, the SA node still sends the impulse down the AV node and Purkinje fibres that spread out across the cardiac musculature and cause them to contract, causing the heart to beat. This will continue for a while till they run out of energy and stop.
You can read more about it here: Wiki

-1 because too generic answer, too copy-pasted and little understanding. Recent research not previewed. To just say that there are that and that pacemaker in heart is not sufficient. More about why -part is needed. Beat itself is not the only process there. There is much more there if you think, for instance, the system in homeostasis. For instance, how are pacemakers kept functional?

@Masi Thanks fr pointing it out. I am sorry you feel that, I look forward to read your opinion in your answer. Secondly, I did not venture into the need for ionic homoeostasis because the question was too generic.

@RoverEye It is not only about ionic homeostasis. The central topic is electrical/mechanical function of the heart together i.e. how heart circulates its energy flow for a sufficient heartbeat. Ionic homeostasis is just a small part of this. Pacemakers are just a small part of electrical system. You do not describe anything about this combination system which the heart contains. There are some good preview articles, for instance, about internal physiology of SA node, describing limitations and targets in this research. Only a few about the integrative system but it still can be described.

For instance, how heart can restore its beating function after i/m KCl in a cardiac surgery where heartbeat must be stop?

@Masi I alwys presumed that after immersing the heart  in KCl during surgery, to restore the functioning of the Na/K pumps you just wash it off with saline? Also, you mention excellent reviews, can you point me to one which you recommend ? And what do you mean by combination system? I have not come across that terminology before, and google is not being helpful today.

KCl is eliminated through normal metabolic processes (kidneys). It cannot be "washed off". The heart is then electrically defibrillated.

@RoverEye One article is The sinoatrial node, a heterogeneous pacemaker structure by Boyett. However, it is not sufficient itself. Much things, I would rewrite completely and extend. Koivumäki et all In Silica ... article is about AF and some introduction for the cell-tissue-organ model in Computational Biology. Clustering, neuronal networks, analysis through different methods and their application - you must understand this from some sources which are outside of Biology. I will let you know when some of better reviews are published in this area.

@Masi If you recommend extended revision, might i suggest you formulate your own answer for this, and that way the OP can have a more complete picture? I'll try and edit as much as can, but you clearly have more to say in this case,, and it appears that you can do it more justice.

Personally, I think this answer is fine... "Does the heart keep beating? Yes, for a while, because it has a built in pacemaker". All the KCI, Kidney, etc (other junk) is good information to have - knowledge is power - but ancillary to actual SIMPLE question. If someone asks how a car engine works, we don't need to learn how brakes make the car stop...

This is a great answer, it just explained what I was looking for

Sydney Ringer showed in 1882 that when the heart, when separated from the body and immersed in lactated Ringer's solution, or even isotonic saline solution, will beat because it has its own pacemaker systems at the level of the Sinoatrial node and Atrioventricular node. This activity will continue until there is insufficient ATP to support it energetically.
The Ringer sources link.
Miller, David J. "Sydney Ringer; physiological saline, calcium and the contraction of the heart." The Journal of physiology 555.3 (2004): 585-587.

I was wondering if a heart keeps beating if it is separated from the body?

Yes, it does. Here's a video showing it (don't click if you're squeamish!): https://www.youtube.com/watch?v=NEuSCPifKhA
It's a classical physiology experiment to remove a heart and keep it beating for some time in a jar with liquid solution. In fact, acetylcholine, the first neurotransmitter to be discovered, was detected via an experiment involving disembodied, still-beating frog hearts. As a result, Otto Loewi (who conducted the experiment) received the Nobel prize.
Of course it won't go on forever, as the heart eventually runs out of nutrients and air (without access to lungs). There are also other problems. Taking steps such as supplying oxygenated blood, putting the heart in an isotonic solution, ensuring stable temperature, adding adrenaline (which stimulates the heart) can extend the duration of beating.

If yes, then why?

Why not? Tissue doesn't die the moment it's removed from the body (it's not like individual cells can telepathically tell they've been cut off, they have no way of knowing). It lives for some time before nutrients are exhausted, blood in capillaries drains off, oxygen is depleted, temperature falls, desiccation occurs, and cells finally die. Note how when an accident severs body parts, they can often be surgically reattached and function normally.
Most tissues don't seem to be doing much when you sever them. For instance, a severed hand won't make any movements (actually severed animal limbs can often keep on twitching) but that is because no signal is coming from the brain. If you could emulate the signal, you could get it to move before necrosis occurs.
The heart has a central cluster of neurons, whose function is to generated a regular signal which causes the heart to beat. Because of this, the heart muscle continues to receive a signal even if cut out. The pacemaker cells don't keep perfect rhythm (and more over heart rate is a complicated matter), so in a normal human the brain fine-tunes the rate of beating through hormones and the vagus nerve. If you cut off the vagus connection, the beating won't be as regular or as rapid as inside the body.

Can you please describe this more closer a central cluster of neurons.

In order to clear some issues with the question and answers I add this one.
The question itself was "will a heart keeps beating if it is separated from the body?"
The answer to this question as it was asked: No, the heart removed from the body "as is" will stop beating almost immediately. Denervation and stopping oxygen and electrolytes supply will cause short fibrillation and eventual asystole.
However, if we convert the question to "What are the conditions that will allow the denervated heart to keep beating?", the answer will be different:
If you supply it with solution of electrolytes it will continue beating for some time. If one want extend that time - other chemical compounds should be added as well as the temperature of the organ should be artificially lowered.
nice link that says exactly the same as my answer (provided by @RoverEye )

I'm sorry mate, but that is not the case. If you remove it completely it still continues to beat for a while. I have seen it first hand, and @Superbest has linked a youtube video showing the same. If we supply it with electrolytes, it will beat longer, but the heart beats without them also.

@RoverEye The heart on the video is partly emerged into the solution. Some hearts will beat more or less, but in general I am sure this answer is more precise. If you were right... people will never "die" if their heart is ok at the death time, their heart will beat indefinitely, but it is not the case.

I am afraid there is a misunderstanding here. The heart is part of the autonomic nervous system and thus has the ability to beat independently of the brain as long as it has oxygen. The heart will eventually stop beating as all bodily systems begin to stop working shortly after brain death. Remember the heart can beat, but your diaphragm and lungs wont. hence the cardiac muscles undergo asphyxiation and die off. However, immediately after death, there is enough oxygenated blood in the body to keep thing moving for a while. A quick google gives enough evidence for this.

@RoverEye the question is about heart "REMOVED from the body" :)

It doesn't matter where the heart is. It can be on a plate for what the heart cares.The heart is a self contained system as far as things go for contraction. As long as it receives electrolytes and most importantly oxygen, it will beat. You might want to read this: http://www.madsci.org/posts/archives/2001-03/986058306.Gb.r.html

Atrial and ventricular fibrillation will continue for maybe 3-5 minutes . 3-5 minutes is a quite a long time.

@RoverEye fibrillation is not beat, you know that very well

True, but its still a muscular contraction. I assumed  from the OP question he meant cessation of contraction, not cessation of beating as we know it. Apologies for the confusion.


text:
GATK workflow for Cancer
I am just starting to learn to use bioinformatics tools.  My university has a limited and expensive bioinformatics team, so I'm mostly on my own except for big questions.
I am planning to use GATK to run 58 cancer control/normal pairs of Exome sequencing data (Illumina) from FASTQ or BAM file format, through the pipeline, with an output of a VCF & MAF format for analysis.
The current GATK pipeline is used for disease but not cancer, so I was wondering if anyone knew if there should be changes made for cancer.  Here's the current pipeline starting with BAM files:

(Non-GATK) Picard Mark Duplicates or Samtools roundup
Indel Realignment (Realigner TargetCreator + Indel Realigner)
Base Quality Score Reacalibration (Base Recalibrator + PrintReads)
HaplotypeCaller
VQSR (VariantRecalibrator and ApplyRecalibrator in SNP and INDEL mode)
Annotation using Oncotator (?)

I'd like some verification that this pipeline will output what I need to run my samples on MuTect, MutSig, or some other analysis program.  I appreciate any advice.

I typically deal with the sequence files upstream of this step, or the VCF files downstream--otherwise I would be more comfortable attempting to provide some helpful feedback. At an extremely superficial level, what you have outlined make sense, but I have just never used GATK firsthand (lots of colleagues have and do).  What about posting this verbatim on these two sites: SeqAnswers, and BioStars? I have seen lots of NGS-related Q&A posted there (your question is essentially a very detailed bioinformatics protocol).

MuTect2 was just released into beta as part of GATK 3.5.  It's based on HaplotypeCaller but makes somatic SNV and INDEL calls.  You can find more information about MuTect2 on the GATK blog and ask any additional questions on the forum.
As a note:  IndelQualityRecalibration is not needed with Mutect2, and there is no VQSR available for somatic calls.
MarkDuplicates -> BQSR -> Mutect2 -> Oncotator is a good basic workflow for somatic variant calling.

This post goes over what MuTec requires as input. mark duplicates and indel realignment will probably have to be done on bam file to use it as input. BQSR is optional and does not change the quality too much. HaplotypeCaller is used for germline not somatic variant calling.
If you have followup bioinformatics questions, you might be able to find answers more quickly on biostars or GATK forums

Thank you!  I didn't know that about Haplotype Caller.  The team at my school suggested it.  I will try biostars or GATK forums.


text:
Consequence of Plants as Incomplete Protein Source
Some years ago, in a 1000~ level biology course we learned that the DNA essentially encodes formulas for creating proteins from amino acids. While the human body can synthesize many many amino acids, some we are unable to synthesize and we dub these as essential amino acids because we must obtain them from our diet. However, surely something must be capable of synthesizing the amino acids, otherwise they would not make it up the food chain to us, which opens the door to the possibility that animals may have varying capabilities in regards to amino acid synthesis. Presumably, this is why cats have to obtain a high protein diet, as they have high need for protein but a poor ability to synthesize them?
Moreover, in doing basic research into nutrition to plan a regular workout schedule there is some need to distinguish protein sources as not all protein sources are considered to be complete proteins because they do not provide all amino acids, or otherwise do not provide them in the right balance.  Particularly, many fruits and vegetables tend to be poor sources of protein.
For plants which are incomplete protein sources, they would seem to lack the ability to synthesize some amino acids. They also would not be able to obtain these through diet, as they're plants. Consequently, does this mean that some plants may not encode for a specific amino acid throughout their entire DNA?

You seem to miss that the concept of essential amino acids is human-centric. Why should a plant encode all the amino acids needed by a human? The plant doesn't need them.

They also would not be able to obtain these through diet, as they're plants. Plant just synthesize carbohydrates by photosynthesis. Plants cannot fix nitrogen and they are dependent on bacteria for that. Though non-parasitic plants can synthesize all amino acids they need, they may not be sufficient for humans as a diet.

Apologies, as my (limited) study was focused entirely on human biology. At the time of study, I had assumed that DNA across species was encoded using the same set of amino acids, and what differentiated them was simply what proteins they made with that given set. But later I considered that this couldn't be correct as a nutritionally incomplete protein source would suggest that a specific amino acid was not being utilized. Broadly, my question was whether the set of utilized amino acids also differs between species.

With a few exceptions among some bacteria, all species on the planet make protein from the same 21 amino acids, and the relative abundances of amino acids is very similar in proteins from plants, animals, fungi and even prokaryotes. See for example this article (available as PDF here). So protein from pretty much any food provides the same amino acids.
The idea that plant proteins are "incomplete" and lack some amino acids seems to be a myth: see for example this correspondence in the American Heart Association's journal Circulation. Plants generally contain less protein than meats per weight, but there is nothing wrong with the protein they contain. Most people in the world obtain the majority of their protein from plants. There are probably some specific parts of some plants that are not nutritionally optimal --- if you for example lived exclusively on peanuts for a year, you might experience some problems ... But any commonsense, varied plant diet will provide all amino acids. A detailed report (and some mythbusting) is here.
Regarding essential amino acids, as mentioned in comments this is a species-specific concept. Humans and most animals can synthesize about half of the 21 amino acids, while the remaining are essential. There are some species differences --- arginine is essential for cats, for example, but not for adult humans. In contrast, plants can synthesize all amino acids. But their protein composition is similar in the end.

So would the amino acid profiles listed on this site (under the protein quality box), for instance, be mostly bunk?

Well, I don't know where that particular data comes from so I can't judge it, but there is certainly a lot of unreliable stuff on various nutrition websites. These companies often want to sell nutrition supplements and tend to have little real data to back up their claims ...  There are certainly some differences between foodstuffs, since we don't eat entire plants but only some parts (see http://ajcn.nutrition.org/content/59/5/1203S.short). But generally, if you eat a varied diet, you will obtain all amino acids you need.

@WYSIWYG Right, I guess that was not very precise. I just meant that they are primary producers (obviously). Will edit.

I changed the 20 to 21 amino acids since most species also code for selenocysteine.

@terdon, selenocysteine may not be so relevant from a nutrition viewpoint since selenoproteins are rare, and the amino acid is probably a very small fraction of total protein. But okay.

@Roland I admit I'm biased since I did my PhD on selenoproteins but it's not that rare. There are 22-25 proteins (depending on how you count them) in the human genome and some animals have many more. They are also essential, those animals that have them can't really survive without them. But yes, I know I'm being pedantic, it's just that it's my field and everyone forgets about poor Sec :)

@terdon Heh, I can sympthasize.


text:
How do I find the number of child terms associated with a specific GO term?
I am trying to figure out how to find the number of child terms associated with a specific Gene Ontology (GO) term using QuickGO, without just counting the amount of terms in a list. For example when I want to find the number of child terms associated with the GO term 'heart development'.
I could just click on the tab 'Child Terms' and count, but what if it would be a very long list, how can I find the correct amount?
I assume there is some way to do this by going to Protein Annotation and applying a filter, but I do not know how to do this.

Any help?

Does it need to be done using QuickGO? Would you be open to downloading the ontology .obo file and parsing it? Do you only want direct child terms or all descendent terms (children, grandchildren etc)?

I am only looking for direct child terms. I would like to find a way to do it in QuickGo if possible, but I am interested in any other methods possible! :-)

I don't think it's possible to do this via QuickGO. You can, however, do it from GO's website, using their GOOSE tool. This offers an interface to the GO SQL database and lets you run queries. It also offers example searches, one of which ("Find descendants of the node 'nucleus'") can easily be modified to return what you are asking for:
SELECT COUNT(*)
FROM
 term
 INNER JOIN graph_path ON (term.id=graph_path.term1_id)
 INNER JOIN term AS descendant ON (descendant.id=graph_path.term2_id)
WHERE term.name='heart development' AND distance = 1 ;

In the query above, I modified the example and changed 'nucleus' to 'heart development' (GO:0007507) and changed distance <> 0 to distance = 1 to make it list only direct descendants. Finally, I also changed SELECT DISTINCT descendant.acc, descendant.name, descendant.term_type to simply SELECT COUNT(*) since all we are interested in is the number of descendants. If you also want their names, leave the SELECT line as is. That will show both the list of names and the total in the output.
The modified query shows us that this term has 30 descendants:

When I also tried running this query in HeidiSQL in UCSC MySQL (database 'go') I found 22 results instead. Any idea why?

@Wolgast different ontology versions presumably. Or a completely different SQL schema. The ontology files are not static, They change quite a bit and quite often. What you need is the number found in the specific ontology file you happen to be using. Don't assume that different online versions will give you the same result.

GOOSE is now deprecated :-(