text:
What is a neuronal loop?
Stanford Medicine's OCD page says that

Many investigators have contributed to the hypothesis that OCD involves dysfunction in a neuronal loop running from the orbital frontal cortex to the cingulate gyrus, striatum (cuadate nucleus and putamen), globus pallidus, thalamus and back to the frontal cortex.

What is a neuronal loop? I have looked it up on DuckDuckGo with no usable results.

The answer is right in the second half of the sentence:

running from the orbital frontal cortex to the cingulate gyrus, striatum (cuadate nucleus and putamen), globus pallidus, thalamus and back to the frontal cortex

Orbital frontal cortex, cingulate gyrus, striatum, globus pallidus, thalamus, and back to (orbital) frontal cortex: these are all different brain regions. If A connects to B, B connects to C, C connects to D, D connects to E, and E connects back to A, you've made a loop.
The page is saying that there is a hypothesis that there is dysfunction somewhere along this loop in OCD; it doesn't say where exactly, but you can infer that changing something along this path would affect all of the other regions connected in the loop in some way, because they are all causally related to one another.

@ds_secret I rejected your suggested edit because the source does not use the word "cause" with respect to this pathway. It may be implied, but I'm not going to put words in their mouth: they are likely using this language intentionally because they have mostly correlational evidence for the relationship rather than causal evidence. Causality is very difficult to show in psychology/psychiatry.

The website states "In the section below, we have outlined a more detailed description of the various theories and hypotheses involved in the biological basis of OCD."

@ds_secret "The biological basis" is not the same as "cause". They are very closely related, may sometimes be treated as "cause", and it's okay if you want to provisionally treat it as evidence for "cause", but I'm not going to use that word.


text:
Does it make sense to express membrane transporters as concentration?
Let's assume a situation where a molecule, S, is transported out of the cell by membrane transporter T. For simplicity we do not consider any other synthesis or production processes. Furthermore, we assume Michaelis-Menten (but another enzyme mechanism would lead to the same question, I think).
Mass balance concentration:
$\frac{d  [S]}{dt} = - [T] \; kcat \; \frac{[S]}{[S] + Km} $
Mass balance amount (moles):
$\frac{d  S}{dt} = - T \; kcat \; \frac{[S]}{[S] + Km} $
In both mass-balances the square brackets [.] indicate concentration.
In order for the units to match, the first mass balance equation expresses the transporter term as concentration (i.e. $\frac{T}{Volume_{cell}}$). Does that make sense? After all, these transporters are stuck in the membrane and not 'floating around' in the cell like the S molecules. Expressing T as concentration assumes that the transporter molecules are equally distributed across the volume, which is not the case. Does this not lead to an overestimation of the reaction rate?
Or did I make a mistake somewhere?

source: Stack Exchange

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

sort: votes


text:
Do larger unicellular eukaryotic organisms have larger organelles?
It seems that many of the giant unicellular eukaryotic organisms (size 1mm and above) are multinucleate but there are some with a single nucleus as well (the genus of Acetabularia). My question is: do they have proportially larger organelles (mitochondria etc.)? If so, can they be seen by a person with sharp eyesight when removed from the cell?

The cell has the amount of organelles necessary to perform its function. If a cell requires a lot of protein, it must have multiple nuclei to produce this protein. If the cell performs many activities that require energy, it needs many mitochondria. So the size and quantity of the organelles will be proportional to their function, not their size. In relation to Acetabularia, it is a single-celled organism visible to the naked eye. The nucleus is visible, but other organelles are not.

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


text:
Is there any known pharmaceutical or chemical that could cause infertility in humans in a single dose with few or no other effects?
Is there any known pharmaceutical or chemical that could cause infertility in humans in a single dose with few or no other effects?
The context is that a number of claims are circulating about vaccination being a "cover" for a plot to render males and/or females infertile. I claimed that it is unlikely that there is a known way to make a person infertile with a single dose of a chemical without significant side effects. However I couldn't find a source.
I'm asking as I believe it is important to have scientifically accurate responses to rumors and the like, even if they are outlandish.

People believing this sort of thing usually aren't that convinced by facts.

@BryanKrause, True, but not-yet-believers may be dissuaded from adopting the belief if contradicting facts are shown in rebuttal to the assertion.

@mgkrebbs Sure, but, given: https://xkcd.com/808/ this is a pretty insane claim. If there were some easy way to infertility with just a pill with no side effects we wouldn't be bothering with vasectomies, tubal ligation, etc. It's a losing battle to argue against things that don't have a basis: the burden of proof has to be on the claim not on the skeptics.

I would be interested to hear from someone with relevant knowledge who can lay out exactly why such a chemical is - or is not - possible. For example has any research been done into chemical alternatives to vasectomies?

yes, chemical castration, or chemical sterilisation! They generally have many side effects. The pill makes millions of female temporarily "infertile" every day. I am not sure how important this question is to the claim though. The simple counterfactual surely is enough? I have had many vaccine and am not infertile. Billions of vaccinations given a year. And in the developed world coverage of upwards of 90% for most vaccines. Wouldn't 90% of the developed world then be infertile?

It's hard to understand why, if such a chemical existed and was available at a price cheap enough to be used in mass vaccinations, it would not be available commercially as a less expensive alternative to vasectomy or tubal ligation.

@reubenmcg but are any of those chemical methods permanently effective in a single dose? My understanding is that they're not?

@jamesqf exactly.

Absence of evidence... years or reading science publications I haven't heard of any. If there was a single dose infertility drug with few side effects this sure would be big, big news. Getting the influential people onside before vaccination would be good, including vaccinating oneself.

". If there was a single dose infertility drug with few side effects this sure would be big, big news. "  -- can you add an answer listing publications where that information would be expected to appear?


text:
Why do meristematic cells have prominent nuclei and dense cytoplasm?
I've been reading about meristematic cells having prominent nuclei and dense cytoplasm. However, I could not understand why it is that way. Could somebody please explain?

Welcome to the site!  "I've been reading..."  would it be possible to elaborate a bit?  Context as to what you have been reading would help us understand the question.  Direct quotes is best with links for reference if possible.  Thanks!

It's from a textbook from school. The book points out the basic characteristics of meristematic cells. "As the cells of this tissue are very active, they have dense cytoplasm, thin cellulose walls and prominent nuclei." I understand that the cells are active but I do not understand as to why that would lead to having prominent nuclei and dense cytoplasm. @rotaredom

Because meristamatic cells are the ones who are undergoing mitosis to increase the height of the plant, and diameter of the plant . They undergo continuous division. So y hey don't mature. They only form the primary cell wall.
I wrote another a answer.  Perhaps this will answer the question

Welcome to Biology.SE! This looks like it could become a good answer, but to do so you need to connect "continuous division" and lack of maturation with the described "prominent nuclei and dense cytoplasm". I would expect it to also mention vacuoles. In addition, answers are much more likely to receive a favorable response if they include supporting references (primary literature is best, but a textbook would work for this). Without that support, your answer is indistinguishable from opinion. ...

... The is a good example of how to format references. ——— You may also want to take the [tour] and then consult the help pages for additional advice on [Answer] effectively on this site. Thank you!


text:
Are these mallards or some other kind of duck?
I stumbled across these ducks in a park in Western Germany. They did not look like mallards to me but this is what image searches and bird identification apps want me to believe. Can anybody confirm that or tell me what other species they are?

Definitely not pure Mallard, as males stand out by their white collars and green heads, females by uniform brown with light flecks. I'm not seeing anything in Google that resembles what you have here - maybe a cross or mutation...? Perhaps a rare breed.

They are hybrid ducks for sure, maybe Brewer's (Mallard x Gadwal) Ducks, common in Europe. Brewer's have high variation in coloring.


text:
Can both lenses of different eyes intentionally focus at different amounts
Can each lens of both eyes have different accommodation rates, or have a different focal length to view objects of different distances. I understand that this can be a problem known as Anisometropia, but can this happen intentionally? For instance, if you place one finger close to the eye but one finger far away, can you focus both eyes in a way so that both appear in view? Assume that the fingers are not placed in the eye's area of visual overlap. I tried this, and I wasn't able to focus on both fingers. But why is this? It seems to be that the eyes would need to be able to individually focus since objects will be different distances from both lenses, like if we look out of the corner of our eye.

It seems plausible to me that chameleons might be able to focus each eye independently, but I wouldn't expect that capability in monkeys and apes (humans).

Welcome Astrovis. Remember to show your own research and where you got stuck as per the [ask] section of the [help] - our requirements.

The answer must be an unequivocal yes and you only need to focus on one object. However, it's not quite how you describe the problem, so not quite what you mean.
If you look at a single object off to one side of your face, but within the "infinity" distance of our eyes (about six meters/19.7 feet), the path lengths between the object and each eye must be different lengths, therefore they must have different focal distances. Obviously the difference isn't massive but the decision to focus on that object is intentional and therefore you have the ability to have different focus points for each eye in an intentional system.
The ability to focus on different objects with each eye is another matter. As far as I am aware, people with Amblyopia (AKA "lazy eye") are not able to focus or not able to focus well with the amblyopic eye and suffer loss of vision control in that eye if not treated. I don't know if anyone has looked at the abilities of people who can independently control their eyes. We have dominant eyes (just as we have dominant hands), so I suspect that we would ignore the input from one eye in a situation where you were looking at two separate objects with two eyes. The Skeptics SE seems to agree with this on a question about pilots using eyes independently.


text:
Export AlphaFold Server file and visualize on PyMol
I am currently working with AlphaFold Server, which is Google DeepMind's public access release of AlphaFold3, a model for biomolecule structure prediction.
I am using it in order to predict protein-protein interactions and I get satisfactory results. However, the Server is limited in its data visualization abilities. For example, I can pan around in the 3D predicted structure, however I cannot zoom in without losing image quality or modify the colors of the proteins (which they had done in their publication in Nature and which you can do in AlphaFold2 or ColabFold).
I believe there is a place where I can export the atomic coordinates and import the files in PyMOL, however I cannot manage to do it. Would anyone more experienced in structural biology and prediction via AlphaFold be able to help me?
I hope this is the right StackExchange to ask this question, hope I have been descriptive enough; and thank you in advance!

When you hit "Download", you get a zip file containing, among other things, atomic coordinates for the five generated models in cif format which can be read in PyMol, Chimera or another program.


text:
What is the least specialized tissue in plants?
According to me, the answer to this question  is meristematic tissue. Meristems have undifferentiated cells and they are totipotent. Even permanent tissues are formed from meristems.
However, while searching this up in the internet, I found out that most of the sites mentioned parenchyma as the least specialized tissue. https://www.toppr.com/ask/question/where-is-the-least-specialized-tissue-located-in-plants/
Although parenchyma is a ground tissue and it perform versatile functions, I don't agree with this.
I would be very grateful to anyone who would clarify my doubt. Thanks.

Why bother defining some tissue as "least specialized"? How would you objectively define something like that?

@Bryan Krause This question has once been asked in a class test.

I think this is a stupid question expecting you to learn by rote memorization some facts given to you in a class rather than having any actual meaning in biology. I'm sorry you're experiencing such a course. Asking for where the least differentiated tissues are would be a reasonable biology question and I'd agree with you that you have the right answer, but it doesn't seem that's the question being asked.

@Bryan Krause Sorry, I don't get why this question is 'stupid' and has no 'actual meaning in biology'. If this question were asked with reference to animals instead of plants, won't it be stem cells?

That would be the other question: asking for the least differentiated tissues would be a reasonable biology question, and yes, for both animals and plants, stem cells, and you might then be asked to identify where those cells exist; in plants, you've identified the meristem. In animals, it would depend on the type of stem cell. But you were asked "least specialized". Maybe in your course they meant "least differentiated" when they asked "least specialized", or else they wanted you to memorize some line in a textbook that said "blah blah parenchyma is the least specialized tissue".


text:
How long does blood stay bright red after being exposed to the air?
I am having a hard time finding the answer to this question. There should be a general time like x-minutes. Does it matter if it is in the sunlight? Maybe even an approximate time like between x-minutes and x-minutes.

Oxygenated haemoglobin is red, deoxygenated haemoglobin is purple (and there is no change in oxidation state when O2 binds to Hb). Oxidized haemoglobin is known as methaemoglobin and is brown in colour.  An example of an oxidising agent that can produce metHb from Hb is ferricyanide.  See 'points to looks for' in this article by Dixon published in Biochemical Education

In addition, blood will clot unless it is collected in the presence of an anti-coagulant such as heparin or EDTA.

I should have added that MetHb (brown) cannot bind oxygen.  In addition, it might be of interest to note that although the binding of O2 to haemoglobin is not an oxidation (we speak of the Hb molecule as being oxygenated), O2 bound to Hb contains no unpaired electrons and (unlike 'free' O2) is not paramagnetic.  To quote from a renowned paper by Pauling and Coryell: "The oxygen molecule undergoes a profound change in electronic structure on combination with hemoglobin"

Blood becomes bright red immediately after the haemoglobin molecules within absorb oxygen from the air, and in air it will maintain this color indefinitely as there is a constant supply of oxygen from the atmosphere and no way to lose it again.

Even when the blood clots?

blood does not stay the same color indefintely it changes as it clots and dries.

I was assuming perfect conditions, like a blood bank.

Geeze, I was not asking how it turns red or why it turns brown. How long does it take for blood to turn brown exposed to the air and assuming that the blood flow has stopped.


text:
What are some good books on oncology?
I'm looking for some book suggestions on oncology, preferably I want them to be fairly recent. I am not worried if they are fairly technical, as long as they have good accurate content and layout.

You could try DeVita, Hellman, and Rosenberg's Cancer: Principles & Practice of Oncology. It has detailed information on both biological and clinical aspects of cancer. Apparently, the publishers regularly release online updates to cover the latest developments in oncology.
Reference to the current edition:
DeVita VT, Rosenberg SA, Lawrence TS, editors. DeVita, Hellman, and Rosenberg's cancer: principles & practice of oncology. 11th Edition. Philadelphia: Wolters Kluwer; 2018. 2432 p.

The must-have textbook: The Biology of Cancer from Robert Weinberg
https://www.routledge.com/The-Biology-of-Cancer/Weinberg-Weinberg/p/book/9780429258794
The Second Edition was published in 2013, I didn't heard about a newer Edition yet but the current one is well recognized in the field


text:
Why do some vaccines lose their efficacy with time?
Why do some vaccines lose with time their efficacy?
The two obvious examples that I have in mind are influenza and tetanus. The former case is clear, as influenza virus undergoes frequent gene reshuffling with similar viruses, which renders the vaccine inefficient against what can be essentially considered a new virus. So the question is about tetanus: does it have to do with the fact that it is bacterial rather than a viral effection? Why the immunity is lost? Are there other similar cases?

@user1136 Here's a study indicating "antibody responses to tetanus declined with an estimated half-life of 14 years (95% confidence interval, 11-17 years)."

@user1136 indeed, I presumed the need for "booster" is because of the declining vaccine efficacy (even if it remains quite high). If you have some experise on this subject, please share with us in an answer.


text:
Is there any limit to asexual reproduction?
As far as I know, DNA degrades with age.  How does this combine with asexual reproduction?
Let's say I have an individual of a plant, and reproduce it asexually.  Once it dies, I reproduce its asexually-reproduced children (or clones, should I say?).  And I (and my descendents) follow that technique throughout centuries.
I assume that since the DNA comes from the very first individual, and it must have degraded over time, at some point it will not be viable to reproduce anymore.
Am I right in that assumption?  For how long can species reproduce asexually without significant degradation?  I guess it also depends on the complexity of the species.

DNA stored in a vial degrades with age as does any biological material, DNA in a living organsim does not.

@John I don't know much about this, so I may very well be wrong, but for a long time I've seen in news and documentaries that DNA gets damaged with age.  For example, https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7846274/.  Does it not get damaged under normal conditions then?

basically any chemical can breakdown with time, biological systems are constantly repairing and/or replacing the chemicals.

So, DNA can fully self-repair to the point that there's no significant degradation over millenniums and tens/hundreds/thousands of generations of asexual reproduction of an individual?

correct, now of course mutations will occur but that is completely diffrent than chemical degridation.

I don't really know a lot about DNA. But I think you meant that the ends of the DNA is "breaking of" while copying it. This is because the protein that is reproducing the DNA needs some time to work rightly. These ends are build by a completely random order of the bases.(the information is saved by the order of them)
At some point important information is breaking of the DNA, that is why we get older.
With every generation somehow these ends somehow have to reset. I have no clue how that is working.
Please correct me if I am wrong. All the information I got from the biology class, so important stuff could be missing. (Or isn't even right)
Anyway I hope I could help you.

Your answer could be improved with additional supporting information. Please [edit] to add further details, such as citations or documentation, so that others can confirm that your answer is correct. You can find more information on how to write good answers in the help center.

Cc: @John  Do you have any comments?


text:
Contribution of ancestors in a person's genome (David Reich's "Who we are and how we got here")
I was reading the book "Who we are and how we got here", written by David Reich. There it has the following explanation:

Females create an average of about forty-five new splices when producing eggs, while males create about twenty-six splices when producing sperm, for a total of about seventy-one new splices per generation. So it is that as we trace each generation back further into the past, a person’s genome is derived from an ever-increasing number of spliced-together ancestral fragments.

Any person’s genome is derived from 47 stretches of DNA corresponding to the chromosomes transmitted by mother and father plus mitochondrial DNA. One generation back, a person’s genome is derived from about 118 (47 plus 71) stretches of DNA transmitted by his or her parents. Two generations back, the number of ancestry stretches of DNA grows to around 189 (47 plus 71 plus another 71) transmitted by four grandparents. Look even further back in time, and the additional increase in ancestral stretches of DNA every generation is rapidly overtaken by the doubling of ancestors. Ten generations back, for example, the number of ancestral stretches of DNA is around 757 but the number of ancestors is 1,024, guaranteeing that each person has several hundred ancestors from whom he or she has received no DNA whatsoever. Twenty generations in the past, the number of ancestors is almost a thousand times greater than the number of ancestral stretches of DNA in a person’s genome, so it is a certainty that each person has not inherited any DNA from the great majority of his or her actual ancestors.

These calculations mean that a person’s genealogy, as reconstructed from historical records, is not the same as his or her genetic inheritance… Yet even if the genealogies are accurate, Queen Elizabeth II of England almost certainly inherited no DNA from William of Normandy, who conquered England in 1066 and who is believed to be her ancestor twenty-four generations back in time.

So, when he talks about how most of the ancestors of a person are not really genetic ancestors, he justifies that at each generation, 71 new splices are generated on average. I got that those splices are the crossovers between mother's and father's chromosomes in a germ cell. So he calculates that a person's genome is derived from the 47 stretches of DNA (46 chromosomes + mtDNA), plus 71 by generation.
To be honest, I just didn't get the logic behind it. The 71 new splices are splices for specific chromosomes, or each splice is a different combination for each of the chromosomes present in the germ cells? And since the splices are combinations of the chromosomes, in any scenario, why would be guaranteed that after 10 generations (1024 ancestors and 757 splices), some ancestors don't contribute to a person's genome? I mean, I can get that the chance is reduced, but I don't get the guarantee, since the stretches are random and uneven, parts of the chromosome could still contain DNA from any ancestor (I think!).
So, to resume:

Considering N generation, why the calculation for where the genome derives would be 71*N + 47?
Considering the point 1 is clarified, why would that be a guarantee that some ancestors don't contribute to a person's genome?

"why would be guaranteed that after 10 generations (1024 ancestors and 757 splices), some ancestors don't contribute to a person's genome?" - Because 1024 is greater than 757? That's the guarantee.

Why couldn't a splice contain the DNA from more than one ancestor? Three, five? Why can we just add up 47 chromossomes + mtDNA with the 71 splices from the previous generation? Do those numbers carry equivalent information? Of course 1024>757, that doesn't add a bit.

If it contained DNA from three people it would be three splices, from five people it would be five splices, that's why the number he comes up with is 757 which is much bigger than 47. How else would you get DNA from more than one person into a stretch of chromosome?

Point #2 is a bit easier to understand (at least to me):
Forget about "stretches of DNA", and focus on individual base pairs. There are around 3 billion (3x10^9) base pairs in a human genome. Each base pair arrived from some ancestor. If we look 1,000 years back, around 40 generations ago (assuming a generation is 25 years), then you have 2^40 ~= 10^12 (1 trillion) ancestors. It's impossible that you inherit at least 1 base pair from each of 1 trillion ancestors, if you only have 3 billion base pairs.
About #1:
You can get an intuition for it by looking at the "extreme case". If you assume that you have only 1 chromosome pair, and there are no recombinations whatsoever, only 2 individuals contributed to your genome: the DNA of only 1 grandmother and 1 grandfather would be present in your genome.
If you allow 1 recombination per generation, then you can have some DNA of all of your grandparents, but you no matter how you recombine the chromosomes, DNA of at least 2 of your great-grandparents will not survive:
Hypothetical stretches of DNA of great-grandparents that end up in next generation. Each letter represents a "DNA fragment":
#1: aaaa
#2: bbbb
#3: cccc
#4: dddd
#5: eeee
#6: ffff
#7: gggg
#8: ffff

Grandparents:
#1+#2: aabb
#3+#4: ccdd
#5+#6: eeff
#7+#7: gghh

Parents:
#1+#2+#3+#4: acdd (we're losing one great-grandparent's DNA here, since only 1 recombination is allowed)
#5+#6+#7+#8: eefh

You:
acdd & eefh (but no traces of "b" and "g" left in your genome)

So by allowing 1 recombination per parent, we got 2 extra individuals possibly contributing to the genome with each generation:

You: max 2 people (your parents)
Including your parents (N = 1): 2 + 2 = 4
Including your grandparents (N = 2): 2 + 2 * 2 = 6

And so on, using 2 + 2*N formula. It works the same way for more recombinations, and more chromosome pairs, but it's a bit more difficult to get the intuition for.
About the "guarantee" that some ancestors didn't contribute to your genome:
The math works out nicely, but of course no one had 1 trillion distinct ancestors 40 generations ago, because there's some intermarrying going on. To say with certainty that:

Yet even if the genealogies are accurate, Queen Elizabeth II of England almost certainly inherited no DNA from William of Normandy, who conquered England in 1066 and who is believed to be her ancestor twenty-four generations back in time.

we would need to know how often royal families intermarry.
If we use 1 recombination per generation, and 1 chromosome pair example, and assume that instead of having 4 distinct great-grandmothers, and 4 distinct great-grandfathers, one has in fact 4 distinct great-grandmothers, but only 2 distinct great-grandfathers, that's only 6 distinct ancestors at N=2. Even though 2 + 2*N < 2^(N+1), one could still inherit some DNA from all ancestors in such a case.

I don't think it's quite right that "no matter how you recombine the chromosomes, DNA of at least 2 of your great-grandparents will not survive." `aaaaaaaa,
bbbbbbbb,
cccccccc,
dddddddd,
eeeeeeee,
ffffffff,
gggggggg,
hhhhhhhh,
|
abbbbbbb,
cccddddd,
eeeeefff,
gggggggh,
|
abcddddd,
eeeeefgh,
|
abcdefgh`

@Darlingtonia Good point. So then you'd also say that the argument from the book would be falsifiable the same way?

The book is correct that as you go back, most of your genealogical ancestors are not genetic ancestors. I find its explanation confusing, but I think it's incorrect to frame this as an absolute certainty rather than a probabilistic certainty. As in my example, the splicing has to occur in just the right way to maximize genetic ancestors. Graham Coop has a series of blog posts around this subject I find helpful. FWIW his modeling doesn't have ancestors dropping out until 5-7 generations back

@Darlingtonia Tricky to wrap my head around it. To me, you more or less proved that the number of genetic ancestors x generations ago being larger than the number of DNA stretches inherited from x generations ago doesn't mean that some ancestors dropped out. That is, the sentence: "Ten generations back, for example, the number of ancestral stretches of DNA is around 757 but the number of ancestors is 1,024, guaranteeing that each person has several hundred ancestors from whom he or she has received no DNA whatsoever." would be falsifiable the same way as my example. Or am I wrong about that?

I think it's that "the number of ancestral stretches" is not a certain number, but rather a probability. There is theoretically a non-zero chance that all ancestors of some generation could pass a piece of DNA to a current individual, but in practice most will be lost because recombination will likely create DNA chunks that exclude some prior ancestors. However, as I said, I personally find the "number of ancestral stretches" to be a confusing way to frame it, so I could be wrong.


text:
Ancestral inference from demographic data
The question is inspired by this one, in the History community:
We consider a population, whereas the data we have the number of migrants arriving every year, as well as the initial and the current population size. We would like to determine what fraction of the population descends from the migrants that came in a particular year.
Naive approach
Let $n_t$ be the number of migrants arriving in year $t$ from the beginning of observation whereas $n_0$ is the number of individuals already present at year $t=0$, $T$ is the number of years. Let $s$ be the rate of increase of the population which we assume constant, so that the number of individuals descending from the immigrants from year $t$ is $$N_t=n_t s^{T-t}.$$ One can now estimate the parameter $s$ from the total size of the population at $t=T$:
$$
\sum_{t=0}^Tn_ts^{T-t}=N,
$$
and we obtain the proportions of interest as $$w_t=\frac{N_t}{N}$$.
Q1: is this a reasonable approach or is there a better way?
Haploid population
The approach above is correct for averages, if we assume haploid population. This is the case when we look only at Y-chromosome, i.e., only at the paternal descent, as suggested in the cited question. There is however demographic stochasticity (or genetic drift - depending on whether we prefer population ecology or population genetics language), which means that the fluctuations in $N_t$ may make our estimates obsolete. I could probably obtain estimates using birth-and-death process ($s=1+b-d$) or a colascent. I am wondering however, if there is a more canonical way of doing it. (this is Q2)
Diploid population
For diploid organism most currently living individuals will have many ancestors from different years. One can thus ask a somewhat different question: how many of the currently living individuals at least one ancestor that arrived in year $t$?
Q3: how could one perform such an estimate?

Since the above seems not to account for different individuals having different numbers of offspring, it doesn't seem well-posed to me. For example, it's certainly possible that limited numbers of individuals gave rise to very high proportions of following generations (I'm thinking of Genghis Khan for example).

@Armand This is what I ask in my Q2. This question can be reformulated as: for how many generations back the estimate is still reliable?

I would say it is never reliable :)

@Armand it is not 100% reliable for sure, but under some conditions the error could be small. E.g., if the population size is constant, we expect that few (and eventually) one genotype will dominate in the long run, but if the number of generations considered is small, the diversity is more or less preserved. If the population is growing we may have even more chances to discerne something.

Humans are well-known for not having uniform family sizes and not choosing mates randomly throughout the population -- that's why any approach that doesn't involve genealogy won't be reliable.

@Armand Genealogy would certainly give better results, but this is not what is given here. I imagine that carrying out a population-wide genetic study in the US is difficult for economic and social reasons.

As mentioned in the OP, one can perform an estimate using birth-and-death process. Let us consider an population from a particular year with the initial size $N_0$ - then the probability that the population has size $N$ obeys the following difference equation:
$$p^N_{t+1} = (1-b-d)p^N_t + b(N-1)p^{N-1}_t + d(N+1)p^{N+1}_t,$$
where $b$ and d are the birth and the death rates (per generation or per year - depending on how we choose the time scale).
Rather than solving for probability, we limit consideration to the mean and the variance:
$$ N_t=\sum_{N=0}^{+\infty}Np^N_t,\\
V_t=\sum_{N=0}^{+\infty}N^2p^N_t-N_t^2.$$
After some algebra we wrrive at:
$$N_t=N_0(1+b-d)^t,\\
V_t=N_0^2\left[(1+2b-2d)^t-(1+b-d)^{2t}\right]+
\frac{b+d}{b-d}N_0\left[(1+2b-2d)^t-(1+b-d)^t\right]
$$
The following analysis is simplified, if we assume slow population change, so that $|b-d|\ll 1$, so that we can switch to continuous time:
$$
N_t=N_0 e^{(b-d)t},\\
V_t= \frac{b+d}{b-d}N_0\left[e^{2(b-d)t}-e^{(b-d)t}\right]
$$
Constant population
Let us first consider the case of a constant total population size $b=d$. The equations above then reduce to
$$N_t=N_0,\\ V_t=N_0(b+d)t.$$
The relative error is then
$$
\frac{\sqrt{V_t}}{N_t}=\sqrt{\frac{(b+d)t}{N_0}}
$$
This error grows with time and eventually becomes large, which is the reflection of the fact that genetic/ecological drift will eventually lead to the extinction or fixation of the allele/population. However, as long as we look at times much shorter than $t\ll N_0/{b+d}$, the error is small and we can make a reliable estimate using the naive approach outlined in the beginning. Indeed, if we take birth/death rate as 1 per generation, and consider the time scales of up to 20 generations (Mayflower came to the US in 1620, 400 years ago) we have the relative error behaving as $\sqrt{40/N_0}$. Mayflower brought about a hundred of passangers, so that the error is relatively big, but for later and much larger imigration waves the error can bemuch smaller.
Growing population
If we assume growing population, $b>d$, and assuming in addition that we look at long time periods, $t\gg|b-d|^{-1}$, we can write the relative error as:
$$
\frac{\sqrt{V_t}}{N_t}=\sqrt{\frac{1}{N_0}\frac{b+d}{b-d}}
$$
This error is small, if $N_0\gg \frac{b+d}{b-d}$. With a birth rate of $3$ children per family, i.e., $b=1.5$ per person, this condition becomes $N_0\gg 5$, i.e., even small immigration waves shoudl be discernable, provided the community grows fast.


text:
Extrapolating rarefaction curves
I am interested in methods for extrapolating rarefaction curves and/or estimating the true species richness. The ecosystem of interest is the gut microbiome and extrapolation is to higher sequencing.
Chao estimator
The ubiquitous approach is the one based on Chao estimator (see Eqs. [7] and [17-19] in this review). However this approach has been recently criticized, since it assumes that the undetected species have the same abundance as the rarest detected species - therefore, at best, it gives only the lower bound for the diversity.
Hill numbers
I am not convinced that using other diversity metrics (most fo which can be represented via Hill numbers) solves the problem. Indeed, most of these metrics have better statistical properties than richness (=number of species), since they give more weight to observe species over the unobserved ones - essentially, they hide the problem under the carpet.
Alternative 1: Using Relative abundance distributions
Making an assumption about actual species abundance distribution (e.g., assuming it to be log-normal or Fisher log-series - whatever is applicable) seems promising. However I have not found any solid publication describing such an approach.
Alternative 2: Comparing rarefaction curves
Alternative approach, employed in package MOMR, is using the rarefaction curves for many samples to extend the other rarefaction cruves.
Question
I am looking for guidance (preferably references with sufficient mathematical details) regarding the two approaches listed above, as well as for possible other alternatives.

Quick question: Are you interested in developing a novel method of estimating true richness, evaluating different methods, or are you simply analyzing your own data and trying to find the best estimates to report? For a community like the gut microbiota, the methods used in sample collection, storage, DNA extraction, sequencing technology, and sequence assembly/curation etc., will probably impact the richness estimates more than the specific diversity metrics used in the final analysis.

@MikeyC I am interested in evaluating different methods. Ultimately, these estimates will serve as a biomarker (one of several) for specific health conditions, so it will be used with extensive amounts of data. For a moment I see that the estimates vary rather wildely... (even though I agree with your assertion that there are plenty of other sources of errors).


text:
Does non-homologous end joining always result in deletion or insertion?
I have been reading about NHEJ and double-strand DNA break repair mechanisms. As far as I understand NHEJ does not use complemantary template, so does that mean when a double strand DNA break is repaired by NHEJ, the breaked DNA could never be repaired to its original form (the form before the double strand break)?

Yes, you are right - NHEJ is working independently of a DNA template, which will in most cases (except for blunt breaks which can directly be joined together) lead to insertions/deletions. In most cases the DNA is different from before. If this leads to problems depends on the affected region of the DNA. Figure 3 from Reference 1 shows the different subpathways of the NHEJ:

Keep in mind that this is an emergency repair trying to save the integrity of the DNA and the cell. There are a few points which show the characteristics as a emergency procedure (see references 2 and 3 for some more details):

Rapid Response: NHEJ is the predominant pathway for repairing DSBs in mammalian cells and is active throughout all phases of the cell cycle, making it a quick response mechanism to DNA damage. This contrasts with homologous recombination (HR), which is primarily active during the S and G2 phases of the cell cycle.

End Resection Independence: Unlike HR, NHEJ does not require extensive end resection of DNA ends before repair can occur. This allows NHEJ to function effectively even when the cell is not actively dividing or when DNA ends are damaged, which is often the case following exposure to ionizing radiation or oxidative stress.

Error-Prone Nature: While NHEJ can efficiently reconnect broken DNA ends, it is also associated with a higher likelihood of errors, such as nucleotide loss or insertions. This mutagenic potential is particularly relevant in situations where precise repair is less critical, allowing cells to survive despite genomic instability.

Backup Mechanism: In cases where other repair pathways (like HR) are compromised, NHEJ serves as a backup mechanism to ensure cell survival. This adaptability highlights its role in maintaining genomic integrity under stress conditions, further reinforcing its classification as an emergency response system.

Although this method is useful in certain situations where DNA damage would lead to the death of the cell, it can cause problems, too. Besides mutations of insertion/deletion of short DNA stretches it can cause chromosomal translocations which may lead to infunctional proteins or the loss of genetic regulation and may cause cancer.
References:

Non-homologous DNA end joining and alternative pathways to
double-strand break repair
Repair of DNA double-strand breaks by mammalian alternative
end-joining pathways
Alternative non-homologous end joining repair: a master regulator of
genomic instability in cancer

Thank you for your answer. Just to be sure, when blunt breaks are joined together doesn't the dna always lose some part of it (e.g deletion) ? So why didn't we just use "in all cases" instead of "in most cases" What I am trying to ask is that is there a way to repair the double strand DNA break perfectly by NHEJ? Some sources says exact repair is possible if the breaks are perfectly compatible, while other dont. I am confused


text:
Comparison of biodiversity between two locations with a known maximum number of species
I would like to compare biodiversity at two sites over a three-year period. We have two plots—one urban and the other peripheral. In both plots, 17 species were initially planted, although with slightly different abundances. Over three years, the number of species and their frequencies were recorded once a year. Species that migrated in were regularly removed, so the maximum number of species is known.
Now, for each site, the goal is to describe how biodiversity developed over the three years. Ideally, both sites should be compared.
I was considering using the Shannon Index and Simpson Evenness for this.
Does that make sense?
Can the two sites even be compared if the initial number of individuals wasn’t the same?
What would you recommend in terms of statistical analysis?
Thank you very much!

So long as you have enough individuals in the sample areas to get some statistical significance out of it, I don't see any problems here. You should have done some a priori test design to work this out before setting up the plots.

Thanks for your respond! Unfortunatly i did not do this priori test before. There are at least 9 Individuals per plot and 4 plots per site. Do you think this could be enough?

This sounds like something your supervisor should be handling. You should look up the Shannon Diversity Index and the Simpson Evenness Index and read about their capabilities and limitations.

You could also use a binomial (or quasi/beta-binomial to account for overdispersion) model for this (how many "successes" are there out of 17 "trials") ?

thanks you. i will think about this.


text:
What kind of spider is this (Colorado)?
I need help identifying what kind of spider this could be. Thanks.

From the leg and body proportions, and the two-tone colouring of the abdomen vs cephalothorax, this is most likely in the Trachelidae family of Broad-faced Sac Spiders.
The Woodlouse Hunter spider (Dysdera crocata) has similar colouring, but the body proportions and, in particular, the chelicerae (mouth parts) are significantly different.
From the Trachelidae family, North America gets the genera Trachelas and Meriola. See https://bugguide.net/node/view/1017586 for additional details.


text:
How to subset SNP data?
I received genomic data from 429 cowpea samples that were sequenced, I would like to know if I can subset this data to select only 110 samples to use for my research. Note that the data is SNP in HapMap format. Please give me some clues. Thank you

We will likely need a lot more information before we can make any suggestions. Why only 110 rather than all? What are the characteristics of the samples and the data? What other data do you have (phenotypes?)? What are some approaches that you've considered? What is your research question?

Not sure what you want to categorise/subset by, but you can definitely subset your HapMap genomic data to select 110 samples.
First, identify the sample IDs you wish to retain. Open the HapMap file in a text editor or use a programming language like Python or R. You can use the data.table or dplyr packages to filter columns corresponding to your selected sample IDs in R. In Python, use pandas to read the file into a DataFrame and select the desired columns. Make sure you include the header and the first few columns (marker information) in your subset. Save the filtered data back to a file in the same format.


text:
Does the hard shell of a chicken egg come from the genetics of the chick or of the mother?
Brought to you by my "what came first, the chicken or the egg?" debate with my elementary aged children.
What causes a chicken egg to develop a hard shell? Does the zygote/embryo create this hard shell, or does the mother's body apply it to the developing chick? In other words, does the hard-shell genotype manifest as a phenotype on the hen or on the egg?
Our hypothetical way to test this hypothesis was, "What would happen if you transplant an egg from a soft-shelled species, say a turtle, into the uterus of a chicken? Would it develop a hard shell?"

A chicken's reproductive system consists of two parts: the ovary and oviduct.

All of the other parts of the egg are added to the yolk as it passes through the oviduct. The chalazae, albumen, shell membranes, and shell are formed around the yolk to make the complete egg, which is then laid. This complete cycle usually requires a little more than 24 hours. About 30 minutes after the egg is laid, another yolk is released and the process repeats itself.

The part of the oviduct that lays down the hard shell is called the uterus, a.k.a. the shell gland.
For an embryo to be present, a chicken's egg needs to be fertilized; a rooster needs to have done his part. Otherwise, there is no embryo, just the mother's genetic material (the egg). This is overwhelmingly the case with store bought eggs; roosters are unnecessary if only eggs are desired. In that case, they're just another mouth to feed (a noisy and disruptive one to boot, but often handsome as all get-out.) Yet unfertilized eggs (containing only the mother's DNA) have a shell.
To test your hypothesis, no transplant is necessary. You could change that to, "What would happen if you transplanted an unfertilized egg into the uterus of a chicken?" Which is, of course, the vast majority of commercial eggs. The shell making "machinery" is all in the mother.
I had chickens, lots of them. I was not willing to kill the young roosters simply because they were male, which meant that I also had an unheard of number of roosters. Well, they weren't exactly unheard. They were incredibly noisy, and if too aggressive, they were the first to become part of a meal. Bless my patient (but not nearby) neighbors!


text:
How can Monstera plant know which leaves to split?
The monstera plant splits some leaves when they put shadow on other leaves behind it, if I understand correctly. But how can the plant know which leave is responsible for the shadowing and therefore which leave to split? Are there other phaenomena similar to this in other plants?

Are you sure it's not just a case of mature leaves always splitting and younger leaves always growing from the bottom? Because you may be mistaking why it splits for how it splits. Many photos show leaves that are split with no obvious shadowing; Just that larger moire mature leaves split automatically.

Plants don't "know" (that I know of) anything, but they do react to many things: hormones, light, temperature, day length, injury, and myriad other factors , many of which I know nothing about. Whatever the case is with Monstera, it's a physiological reaction to some stimulus, not an intellectual one. If I had a few hours, I could give you a layperson answer based on this paper, but I have fewer hours left than you probably do, so have at it!

Thanks for the hint. Obviously, I am looking for a mechanism, yes.


text:
Location of Kozak Sequences
Are Kozak sequences always found within the first exon? I know that the first exon must be right next to the promoter so that RNA polymerase can start transcription, but just wondering how that would work if kozak sequences aren't always found in the first exon.

Welcome to the site. Please take a [tour] and visit the [help] for more information on this site. Please also note that one of the requirements for this site is some prior research by you into what might be the answer to your question. On that note - what comprises a Kozak sequence (in bases) and how does this answer your question...?


text:
How do you calculate the time until the steady-state of a drug?
I encountered a mathematically intriguing conundrum, in that it's related to medicine but is centered around mathematics.
Suppose drug A has a half-life in the body of 30 hours. The patient takes 40mg once per day. How long is it then until the patient then reaches steady-state dosage in the body (within some tolerance interval)?
The answer should be about one week, but I'm trying to understand the math behind "why".

Here's a short simulation that show the process. You can find more about how to simulate a half decay process here
You can notice that the first dose of 40 mg should decay to 20 mg in 30 hours, but the process is cut by the new dose.
The daily oscillation reaches a steady state around the seventh day (you can find small changes in the data, but not appreciable at the naked eye)

import matplotlib.pylab as plt
import numpy as np

ndays = 12
ttot = 24*ndays
dt = 0.25

time = np.arange(0, ttot, dt)
Concentration = np.zeros(len(time))

conc = 0
tau = 30./np.log(2)  ## exponential decay time constant from half-life value
dosis = 40

for k, t in enumerate(time):
    if t%24 == 0:   ## If the time is multiple of 24 hours, the dosis is given
        conc += dosis
    Concentration[k] = conc
    ### update concentration decay
    conc = conc*(1 - dt/tau)

ax = plt.subplot(111)
ax.plot(time, Concentration)
ax.set_xticks(np.arange(0, ndays * 24, 24 ))
ax.set_xticklabels(['%d'%d for d in np.arange(ndays)])
ax.set_xlabel('Days')
ax.set_ylabel('Dosis remaining (AU)')
plt.savefig('Steady_state.png', dpi = 300)

Can you explain where the 0.693 in the calculation of tau comes from?

0.693 = ln 2. Assumed exponential kinetics for half-life

you could use np.log(2) to make that clearer ...

Thanks for the nice plot. Is there a way you could simulate what the effect would be if the daily dosage was instead an "average" over a 24 hour period? That should give you a nice differential equation you can solve explicitly.

Glad you like it. The differential equation you propose is the one in #thegreatemu answer. In that case D = dose/24 (mg/hour). If you simulate it, the dynamic is the same he found analytically. The Euler step can go like this "conc = conc*(1 - dt/tau) + dt*dosis/24"

heracho's answer does a good job showing the realistic timing. However you can do a pretty good approximation with a simple differential equation, where you approximate the daily dose as a continuous input. If $C(t)$ is the concentration of the drug at time $t$, then
$$ \frac{dC}{dt} = D - \frac{\ln{2}}{t_{1/s}} C$$
where $D$ is the dose rate, 40 mg/day. The steady-state condition is reached when $\frac{dC}{dt} \to 0$, which tells you that the (average) steady state concentration is $$C_{ss} = D\frac{t_{1/2}}{\ln{2}} \approx 72.13\mathrm{mg}$$ which agrees pretty well with heracho's plot.
The full solution to the differential equation, assuming $C(0)=0$ is
$$ C(t) = C_{ss}\cdot(1-\mathrm{e}^{-t\ln{2}/t_{1/2}})$$
From here you can see clearly the answer to the "when do we reach steady state?" question is driven by the exponential term, which is negligible by ~5 half-lives.
Having stolen @heracho's plotting script and added the following lines:
conc_ss = dosis*(tau/24)
concavg = conc_ss*(1-np.exp(-time/tau))
...
ax.plot(time, concavg, label='continuous dose approximation')

you can see that this approximation follows the mean of the true concentration pretty well after just a few half-lives.

Very nice, I appreciate the simpler differential equation that matches closely, it's a bit easier to analyze than something that oscillates up and down so much.

Let's take a moment to think about what it means for a drug dosed multiple times to reach "steady-state": "steady-state" would mean that it doesn't matter much if you've been taking the drug for N or N-1 days, if you've reached "steady-state" the concentration of drug is steady from day to day (of course, with a 30-hour half-life, it's not going to be particularly stable within a day, assuming relatively rapid uptake).
It's also important to recognize that simple exponential decay is a scenario that occurs when the fates of particles are independent of each other. Therefore, one way we can think about comparing day N to day N-1 would be to consider how much drug is still around on day N from the first dose, because that's the only difference in the history of doses.
For a drug with a 30-hour half-life, after 7 days you could say 7 days * 24 hours/day * 1 half-life/30 hours = 5.6 half-lives have passed.
After 5.6 half-lives, you could say 0.5^5.6 = about 0.02 or 2% of a drug is remaining relative to the original concentration.
Another way to say that is that the difference at the end of the 7th day between having had the drug for 7 days vs. having had it for 6 days would be that 2% remaining from day 1. That seems roughly reasonable to consider a "steady-state".
This is all assuming simple exponential kinetics, other assumptions will change the nature of the problem. It's also important to recognize that for a drug with a 30-hour half-life given once daily, the concentration will never really be "steady" but will still have a clear oscillation after each dosage, depending on the kinetics of release/uptake.

This seems like a good start, but it could be more clearly framed in terms of build-up of concentration over time. The time scales are the same for loss from steady-state vs. build-up of a monomolecular/saturating curve starting from zero (basic dynamics are $1-\exp(-kt)$), so a half-life describes the time taken to decrease the distance from the current concentration to the steady-state concentration (i.e. after 5.6 half-lives we are within 2% of the steady state).

@BenBolker I was trying to go from a simpler intuitive approach to exponential decay, which is to recognize that every molecule is independent of the others. That means that on the 7th day you can know how much of the first dose is still present, which tells you whether you should care that you're on the 6th versus 7th versus 8th day of dosing. Once it doesn't matter, you can effectively consider that "steady-state", though of course there is still some accumulation forever, we just have to decide at what point we don't care anymore.

This is a very clever approach. To just check how much of the first dose is still in the body at day N, and compare the difference of having taking it or not (N v/s N-1 days).

The first two paragraphs had me thinking this answer had missed the point of multiple dosing, but now I see it's a very elegant solution to the problem (avoiding the work of simulating an ODE with a series of impulses).

@electronpusher I think I probably could have done a better job of setting up the conceptual approach. I'll try to improve it.

I like @thegreatmu's answer, but I think we can get closer to the original
problem.
During the day, the concentration decays according to
$$
\frac{\partial C}{\partial t} = - \frac{\log(2)}{t_{1/2}} C
$$
pretty much like @thegreatmu's equation, but without the term $D$, because
during the day, no medicine is added.
At a fixed point during every day, say at breakfast, the medicine is ingested,
and the concentration instantly increases by the dose $D$. At such moments,
whe have both the low concentration $C_-$ and the high concentration $C_+ = C_- + D$.
Let us measure time in days (the half-time is 1.25 days), and let us take breakfast-time to be the moments $t=0$, $t=1$, $t=2$, etcetera.
The concentration at any time $n-1\leq t < n$ in the $n$-th day is given by
$$
C(t) = \exp\left(- \frac{\log(2) }{t_{1/2}}  (t-n+1)\right) C_+(n),
$$
so just before taking the medicine again, we have the low concentration
$$
C_-(n+1) = \exp\left(- \frac{\log(2) }{t_{1/2}} \right) C_+(n).
$$
and the high concentration is given by
$$
C_+(n+1) = D + \exp\left(- \frac{\log(2) }{t_{1/2}} \right) C_+(n).
$$
From here it is pssible to calculate the high concentration for any day $n$:
\begin{eqnarray}
C_+(0) = D &,&
C_+(1) = D\left(1 + \exp\left(- \frac{\log(2) }{t_{1/2}} \right)\right), \cdots
\nonumber
\end{eqnarray}
leading to
\begin{eqnarray}
C_+(n) &=& D\sum_{j=0}^{n} \exp\left(- \frac{\log(2) }{t_{1/2}} \right)^j
=
    D\frac{1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)^{n+1}}
          {1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)}
\nonumber
\end{eqnarray}
and at $n-1\leq t < n$, any moment in the $n$-th day, the concentration is given by
\begin{eqnarray}
C(t) &=&
    D\frac{1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)^{n}}
          {1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)}
\exp\left(- \frac{\log(2) }{t_{1/2}} (t-n+1)\right)
\nonumber
\end{eqnarray}
we can compare the current concentration $C(t)$ to the concentration
$C(t+m)$, the same time of day, but $m$ days later:
\begin{eqnarray}
C(t) =
    \frac{1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)^{n}}
         {1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)^{n+m}} C(t+m)
=
    \left(1-\exp\left(- \frac{\log(2) }{t_{1/2}} \right)^{n}\right)C(t+\infty),
\nonumber
\end{eqnarray}
where $C(t+\infty)$ means 'at this time of day, in steady state'.
Therefore, we can say that the concentration is in steady state when
\begin{eqnarray}
    \exp\left(- \frac{\log(2) }{t_{1/2}} \right)^{n} < \epsilon,
\nonumber
\end{eqnarray}
where $\epsilon$ is small, for instance $\epsilon = 0.01$ when
you consider 'in steady state' to mean 'less that one percent away from steady state'. This will be after $n$ days, where $n$ is given by
\begin{eqnarray}
    n >  {}^2\log(1/\epsilon)~t_{1/2}.
\nonumber
\end{eqnarray}
In the case where the half-time is $1.25$ days, and $\epsilon=0.01$, we get
$n > 8.3$: just over a week. After 7 days, steady stage is approached up to
2%.


text:
Is it true that every organ was inherited from common ancestor?
For me it seems nearly impossible that two different branches of evolution came up to same organ or feature.
Is there any proof or refutation that every organ or feature was inherited from common ancestor?
Is there any mechanism of adopting organs or features from different branches?
Example of organs and features:

Eyes
Body symmetry
Head
Lungs

This question seems relative a bit: What is the evidence that plants and animals had a common ancestor?

Eyes are a great example of convergent evolution: https://en.wikipedia.org/wiki/Convergent_evolution Even if you don't believe it, this is what happened.

The catch-all terms in your list belies the differences that exist.  There are radically different kinds of eyes. For example, your eye, insect eyes, octopus eyes, and  lobster eyes are all wildly different. The same is true with lungs (invertebrate lungs vs vertebrate lungs). This is also true with hearts, limbs, ears, mouthparts, and taste/smell organs

"Is there any mechanism of adopting organs or features from different branches?" We suspect this occurred on a microrganism level where organisms can absorb each other and organelles, but not a between larger multi-celullular organisms.  However, that kind of stuff tends to go so far back that they might not have been considered different branches at the time.

For me it seems nearly impossible to imagine the changes possible over millions of years. A dollar is a very small thing, but think how different your life would be with 50o million of them. Now think how much greater a year is than a dollar. It doesn't matter what seems impossible to you or to me. It is what it is.

what prior research have you done, eyes and symmetry have evolved independently several times. eyes are particularly famous for it since it has happened almost a dozen times. also there are no common organs among animals and plants are you perhaps confusing organelles and organs?

@DKNguyen but could all these different eyes be evolutionized version of the initial primitive eye?

@k06a See answer.

It seems that some features/organs could appear during evolution independently for different branches of evolution.
https://www.scienceworld.ca/stories/evolution-flight/

Flight appears to have evolved separately four times in history: in insects, bats, birds and pterosaurs. These four groups of flying animals didn’t evolve from a single, flying ancestor. Instead, they all evolved the ability to fly from separate ancestors that couldn’t fly. This makes flight a case of convergent evolution.

For chordates and, more specifically, vertebrates, most organs do share a common ancestry. Some organs with different functions share the same origin too. For example: cephalochordates have a iodine mucus-producing gland called endostyle, which roots back to the first protochordates. Its function is to capture microorganism and detritus.  In vertebrates the endostyle became the thyroid gland, which obviously has a different function but still secretes iodine.
However, if we take a look at Metazoa in general (=basically all animals grouped into one taxon) some organs definitely have different origins because of convergent evolution. For example, vertebrates and cephalopods both share camera-type eye and a similar central nervous system but those features evolved separately. Same thing applies for arthropod hearts and cardiovascular system compared to vertebrates’ one. Both have a closed circulatory system but that’s a certain case of convergent adaptation, since deuterostomes’ and protostomes’ last common ancestor was an acoelomate and didn’t even have a cardiovascular system in the first place.

I'm impressed there is evidence for so fundamental an necessary an internal organ as the heart, What is the evidence for complete independence for vertebrate and cephalopod and vertebrate eyes? Is there any? The eyes obviously developed along different lines but from context of OP's question, it seems even originating from a common light sensitive organelle in a micro-organism could disqualify it as being of truly independent origin.

"For me it seems nearly impossible that two different branches of evolution came up to same organ or feature."
Depending on what level of organization you are talking about ("feature" is rather imprecise), the answer differs somewhat, but in general it is quite possible and instances are known, as laid out a bit in other answers.
If you are talking about an "organ", depending on what you mean, the panda's thumb might be an example, or eyes.
What's instructive about the example of e.g. cephalopod vs. vertebrate vs. insect eyes is that while they are quite different structures, they are all actually quite similar at the molecular level, for example using homologous opsin proteins, chemical processes, and in fact arising from the same ancestral tissue lineage. The case is instructive in that the evolution was parallel, arising from a shared set of input components among all lineages, rather than convergent, arising in a totally de novo fashion (though in general people seem to prefer "convergence" as the more general term). This kind of parallel evolution is much more frequent than completely independent convergent evolution.
More generally, convergent evolution occurs more frequently between closely related lineages than between distantly related lineages, arguing that the shared evolutionary context makes it more likely that the same adaptations can occur. Similarly, such convergence tends to reuse the same molecular components rather than to derive similar components independently.

The catch-all terms in your list belies the differences that exist. There are radically different kinds of eyes. For example, your eye, insect eyes, octopus eyes, and lobster eyes are all wildly different. The same is true with lungs (invertebrate lungs vs vertebrate lungs). This is also true with hearts, limbs, ears, mouthparts, and taste/smell organs
For the eye, specifically, it would not be surprising if we were unsure whether it can be evolutionized to a single initial version of a primitive eye. The issue is that if you go far back enough we just aren't sure due to lack of fossil evidence. The simpler things are, the less variation there can be and the more likely two independent things will end up the same. If there were a common origin for all eyes, it would likely be a light sensitive organelle on the microorganism level and you can see why fossil evidence for such a thing is tricky to discern even if found it.
Even going as far back as the mysterious Pre-Cambrian may be insufficient.  Jellyfish, molluscs, and worms (our ancestors) already existed, and likely arthropods (marks that look like leg tracks have been found), all of which are known to be capable of light sensitivity...in modern times at least.
But let's not forget plants can detect light too and they go back even farther so that could be your evidence for convergent evolution of eyes...but again, it's also  not a sure thing unless you can find and compare genetic evidence for light sensitivity between living plants and animals).
But eyes are a difficult and unnecessary example if you just want completely independent convergent evolution examples. Legs are dead easy. Arthropods and "fish" diverged before the "fish" even had fins (from which limbs would evolve for all tetrapods), and arthropods already had legs.
Another easy one is wings between insects, bats, pterosaurs, and birds. Less "optional" structures such as internal organs (and especially body symmetry) tend to go back really far and make things difficult.

It is true as a general rule, with exceptions.


text:
Why is scaffolding necessary in organ printing?
Most organ printing techniques seem to use a structural element for newly created cells to anchor themselves to, be it a biocompatible plastic or some natural material. Naively, I would assume that organs can be generated just the same as they are in an embryo, so why is scaffolding necessary? Here are my guesses:

The time taken to grow an organ to an adult size this way would take too long to be practical
Making the necessary germlines to emulate those formed in gastrulation is to difficult
Organs grown this way collapse into themselves without the exact environment present in an embryo (the pressure in an embryo could be necessary, or the formation of all organs concurrently allows each particular organ to maintain a solid boundary between itself and its neighbors)
Without scaffolding, organs printed in vitro would not generate the auxiliary structures necessary to be useful (nerve endings, blood vessel, etc.)

You can't extract the print a heart "code" from DNA, that's not how DNA or development works. Each organ develops according to a pattern influenced by all the other surrounding tissues. It's quite an intricate dance, with folds and migrations occurring at specific times. That's why you can't regrow lost digits or limbs, or replace damaged brain tissue: the connections those pieces make with the rest of the body are only possible during a specific time in development when all the correct cues and relationships are there.
In other words, to generate an organ the way it is generated in an embryo, you need a whole embryo. To generate one to an adult size that way, you need to grow a whole adult. Science fiction has used this as a plausible approach, but real society finds it problematic to grow people for the sole purpose of organ harvesting.
Read, for example, the Wikipedia article on heart development:

In the splanchnopleuric mesenchyme on either side of the neural plate, a horseshoe-shaped area develops as the cardiogenic region. This has formed from cardiac myoblasts and blood islands as forerunners of blood cells and vessels.[5] By day 19, an endocardial tube begins to develop in each side of this region. These two tubes grow and by the third week have converged towards each other to merge, using programmed cell death to form a single tube, the tubular heart.[6]
From splanchnopleuric mesenchyme, the cardiogenic region develops cranially and laterally to the neural plate. In this area, two separate angiogenic cell clusters form on either side and coalesce to form the endocardial tubes. As embryonic folding starts, the two endocardial tubes are pushed into the thoracic cavity, where they begin to fuse together, and this is completed at about 22 days.[7][2]

The only place you get a splanchnopleuric mesenchyme to start with is going to be a developing embryo. The heart is forming from different clusters of cells migrating and differentiating in a particular pattern.
Scaffolding is more of a trick. It gets you away from the organ-level patterning that shapes the entire structure and lets you deal with just the functional units: individual cell types like myocytes that behave sufficiently robotically that if you create approximately the right conditions they will settle in to do their job.


text:
How does blood flow to and from the brain when a choke is applied to one side of the neck
In the sport of Brazilian Jiu-Jitsu and Submission wrestling there are some chokes that only target one side of the neck, meaning only one of the carotid arteries. In the context of blood flow, how is this possible since we have two carotid arteries located on the "right" and "left" side of the neck?
A friend of mine gave an explanation that the neck, head and chest can be seen as a circular flow of blood and if one where to stop the flow in the left or right part of the neck than that would stop the "circular flow". Is this correct?
Here is a video showing an arm in guillotine which I believe (and the author states) only target one of the carotid arteries: video
To concisely summarize the question: How does a single sided neck choke work in the context of blood flow?

I do not know what happen in single side choke . But definitely there is circular flow of blood at the base of brain called circle of willis that connects two internal carotid arteries coming anteriorly and a basilar artery that is formed by joining of two vertebral arteries of both side. It can help to flow the blood coming through carotid artery of other side to the choked side.

Your question is regarding what happens when one temporarily compresses a carotid artery on one side.
The key point here is whether the brain can receive adequate blood supply in that time period. The general answer (for a healthy individual with standard anatomy) is yes.
Put simply, the blood supply to the brain comes from two sources:

internal carotids - supply the front of the brain (via ACA and MCA) on their respective sides
vertebral arteries - join to form basilar artery, which supplies the back of the brain and brainstem/cerebellum

These are all joined together in a circle of Willis (via communicating arteries), which forms a clever network of collateral connections that can be rapidly activated in case of sudden interruption of flow on one side (primarily in the case of unilateral carotid occlusion). This is thought to be deployed in a matter of seconds.
In addition to this, though less relevant in this scenario, there is also a secondary network of collateral blood supply, coming from leptomeningeal and ophthalmic arteries (but they are thought to be more important on longer timescales, on the order of hours or more).
Of note, there are significant variations in how different blood vessel networks are connected (within the circle of Willis and otherwise) among different individuals, with up to 50% having some anatomical variation. This means that certain adaptations in blood flow may not be possible (and that some individuals may be much more susceptible to adverse effects).
Outside of martial arts and in the world of medicine, the following scenarios are important:

carotid massage (a technique for treating certain heart arrhythmias/SVTs) - is classically taught to be performed on one side at any given time, due to risk of causing fainting if performed on both carotids simultaneously
carotid stenosis - this is part of the spectrum of manifestations of vascular disease/atherosclerosis, where plaque deposition partially (or completely) blocks the lumen of the artery. It can be completely asymptomatic (sometimes even in near-complete occlusion) or it can lead to devastating strokes. This, again, highlights that the blood supply to the brain can be somewhat of a mystery (and that individual collateral networks and anatomy matter).

Finally, a word of caution: carotid manipulations (medical or otherwise) do not come without risks. In particular, they are contraindicated in individuals with heart arrhythmias (as carotid massage can potentially precipitate these), history of stroke/heart attack and anyone who might have or is known to have carotid stenosis (as the last thing you want is to dislodge the atheromatous plaque and send it flying into the brain to cause a massive stroke...).

Some sources / further reading:
Romero, José R., et al. "Cerebral collateral circulation in carotid artery disease." Current cardiology reviews 5.4 (2009): 279-288.
Thanvi B, Robinson T. Complete occlusion of extracranial internal carotid artery: clinical features, pathophysiology, diagnosis and management. Postgraduate medical journal. 2007 Feb;83(976):95-9.
Maguida, Gino, and Ashfaq Shuaib. "Collateral circulation in ischemic stroke: an updated review." Journal of Stroke 25.2 (2023): 179-198.


text:
How to measure cytochrome P450 activity in owls?
I hope you're all doing well. I'm currently working on a research project focused on owls and their exposure to anticoagulation rodenticides such as brodifacoum. As part of my study, I need to measure cytochrome P450 enzymes activity in these birds to assess their ability to metabolize these toxins.
I'm curious to know if there are assay kits or ELISA kits commonly used for measuring cytochrome P450 activity in owls. Any insights or advice would be greatly appreciated. If there are alternative methods or techniques that may be useful in this context, I am open to exploring them as well.
Thank you in advance for your help, and I look forward to any insights you can provide.


text:
When does oxidation destroy prions?
It seems like a no-brainer than oxidation, playing the, er, role it does in the universe, would destroy prions just like it destroys everything else.
But when does it do that? I assume this has been studied, since people handle them in laboratories and may need to sterilize their instruments.

Are you asking how strong of an oxidizing agent you would need to eliminate any infectious prions?

Rust is not destroyed by oxydation :P

Just to add to my previous comment: what I meant is that "oxydation destroys everything" is a gross misunderstanding. Oxydation, just like reduction, transforms one molecule into another. Oxydising an alcohol you can get a carboxylic acid. Have you "destroyed" the alcohol? Sure, but you have also formed a carboxylic acid!

Prions are fairly resistant to traditional method of sterilization.
Citing from: Guideline for Disinfection and Sterilization of Prion-Contaminated Medical Instruments

The prions that cause CJD and other TSEs exhibit an unusual resistance to conventional chemical and physical decontamination methods. Because the CJD agent is not readily inactivated by means of conventional disinfection and sterilization procedures and because of the invariably fatal outcome of CJD, the procedures for disinfection and sterilization of the CJD prion have been both cautious and controversial for many years.

The same paper also lists several effective and ineffective methods for sterilization (efficient method have > 3log10 reduction in prion content within 1 hour)
I am not going to list them all here but you can see Table 2 in the paper.
Just to answer your specific question about oxydation, certain oxydising agents are effective, others are not. Similarly certain autoclaving (=sterilising with heat) procedures are fine and others are not.
Another interesting paper Methods to Minimize the Risks of Creutzfeldt-Jakob Disease Transmission by Surgical Procedures: Where to Set the Standard?

If you google "prion oxidation" you'll see that there has been some research into this over the last 10 years, though it seems to be a minor focus of the overall field. There seems to be some interest currently in using ozone to inactivate prions. One company that offers a ozone-based sterilizer claims to be testing for effectiveness against prions, but that is an extremely long-term process (you have to isolate prions, subject them to your sterilization process, then inject them into the brains of mice or hamsters and wait to see if disease develops).
If you're curious as to how researchers sterilize material in the lab, I may be able to shed some light on that (I worked in a prion lab in 2006-07). Generally, disposable instruments and materials are used whenever possible and incinerated after use. Glassware is submerged in acidic detergent and autoclaved for 4 hours at (I believe) 132C. As far as I know, there is still no known way to completely sterilize stainless steel instruments - this was being studied at the time in the lab where I worked. Any piece of equipment that came in contact with human or bovine prions was kept in Biosafety Level 3 facilities permanently. I'm not sure what the decontamination process was for retired equipment. Separate equipment was maintained for "non-human pathogen" prions - hamster, mouse, etc.

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


text:
Can oxygen diffuse out through the lungs?
Let's say you accidentally walk into a room pressurized with pure nitrogen (or you're jettisoned into space). Within a couple of seconds, the partial pressure of oxygen within your lungs drops to 0. But there's still a (small?) reserve within your bloodstream.
Now, will the oxygenated blood travel to your lungs? If so, will the usual gas exchange reverse, with oxygen going from your blood to the oxygen-poor environment?

Yes.  Blood will continue to circulate (as long as you're alive and your heart is beating), so will eventually come in contact with your lungs.  Once there, the gasses (O2, CO2) will cross out into the lung and be exhaled.  All diffusion is driven down a concentration gradient.

Okay, two questions I think need to be explicitly asked: (1) In my basic research I read here that something goes on with 2,3-BPG near the heart which allows deoxygenated RBCs to be separated from the oxygenated ones – does that play a role here? (2) Can the concentration gradient just flat-out strip the oxygen from hemoglobin?

There are many factors - BPG concentration, hydronium concentration (i.e., pH), cooperativity - that modulate the actual Kd for oxygen of hemoglobin (described here http://employees.csbsju.edu/hjakubowski/classes/ch331/bind/olbindhemoglobin.html as well as other places), but the physics/chemistry is unavoidable: if one moves to an atmosphere devoid of oxygen, oxygen will leech out of the blood.  http://www.olemiss.edu/depts/chemistry/courses/chem471_99/L12_6.pdfhas some graphs that show binding as a function of pO2.

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


text:
What makes a DNA sequence motif a DNA sequence motif?
I've read about DNA sequence motifs, but still don't understand what makes some sequence a DNA sequence motif. Is this a well-defined concept? For, example,
What are DNA sequence motifs? says:

Sequence motifs are short, recurring patterns in DNA that are presumed
  to have a biological function. Often they indicate sequence-specific
  binding sites for proteins such as nucleases and transcription factors
  (TF). Others are involved in important processes at the RNA level,
  including ribosome binding, mRNA processing (splicing, editing,
  polyadenylation) and transcription termination.

For example, I've been working with Recombination Signal Sequences (RSS), which are important in immunology. However, from what I can tell, RSS are not themselves motifs, but they do contain motifs. E.g. Conservation of sequence in recombination signal sequence spacers
says

Previously, the RSS has been described as possessing both a conserved heptamer and a conserved nonamer motif.

So, apparently, the heptamer and nonamer component of the RSS are themselves motifs, but the RSS itself is not. Can anyone explain why? Thanks.

Motif means a pattern. For any array of objects to be called a pattern, it has to have at least more than one instance.
To define more precisely, a motif should have significantly (statistical) higher occurrence in a given array compared to what you would obtain in a randomized array of same components (expecting at random).
[Here i use the term array to mean an arranged set such as a genome (array of nucleotides with their position in the sequence) or a protein structure (array of amino acids with their vector positions). A randomized array with same components is generated by shuffling the elements and their values; for e.g. assign the same collection of nucleotides to random positions]
RSS nonamers/heptamers are called a motif because they are a pattern of sequence which a) has more than one occurrence b) the occurrence is significantly higher than a random expectation. Generally in sequence motifs randomization is not usually done because the genome size is too huge. In such conditions if your sequence has >= 2 occurrences throughout the genome it may be considered a motif. Lets assume a 10bp sequence. Its random probability = 1/4¹⁰. Therefore in 1048576 bp (1Gb) there is a likelihood of finding one instance of that sequence on random. So I assumed that if that sequence has >=2 instances it can be called a motif. However, I am not sure about this.
RSS is not a motif because the entire RSS is not conserved; the conserved 'motifs' are interrupted by sequences. This is analogous to lets say a promoter which is a functional region of the DNA. Promoter can have binding sites for different transcription factors and these operator sites (TFBS) can constitute a motif (if they are conserved) but not the entire promoter.

Thanks for the reply. I've read such definitions before, but they have never seemed very precise to me. Does this mean, for example, that the RSS cannot be determined to have "significantly (statistical) higher occurrence in a given array compared to what you would obtain in a randomized array of same components (expecting at random)."?

Edited the answer.. see

I'm confused. So, you are saying that all of the RSS and the included heptamer and nonamer are motifs? I had the impression that the RSS contains motifs but are not themselves motifs.


text:
Water transport to the Shoot apical meristem in a seedling
I am familiar with how plants transport water from the roots to leaves. Specifically, Water evaporates from the leaf surface, generating a negative pressure gradient which pulls water up from the root.
What is unclear to me, is how water is transported to the shoot apical meristem of a seedling, where there is no leaf above it (see red arrow on image attached).
Are there stomata on the shootapical meristem from which the water can evaporate? Or is there some other mechanism which transports water up here? Thanks.


text:
Does Lymph Clot?
According to my professor, lymph has the ability to clot because it contains plasma which has dissolved fibrinogen. He says that as long fibrinogen is present in a bodily fluid, it should be able to clot.
I always thought lymph would not be able to clot as it lacks the thrombocytes to initiate the coagulation pathway. Without thrombocytes the Stuart–Prower Factor would never be released into the lymph stream, and hence the coagulation cascade never occurs.
So does lymph have the ability to clot, and what would be the mechanism for the clotting of lymph?

There happens to be a relatively recent (2021) review on just this subject, also known as lymphatic thrombosis. The text is open access. Unfortunately I don't have time to write a full answer now (hence the comment), but if you're having trouble understanding the paper let me know and I'll write an answer covering the areas you have questions about.


text:
Insect identification request : is it any species from muscidae family or some another?
I belong to Haryana , India
Sudden I found this type of insect in my classroom
and it stung me suddenly because ,I felt itching in my hand for about 15 mins , so please tell me, is it poisonous or not and which type of Arthropod is it ?
Characteristics:-
Two horn like protrusions on its head ,
A stingy tail,
Eyes are not clearly visible,
Winged ,
1-1.5cm in size
(Sorry for blurred image )

The photo is far to unclear to be able to tell anything other than "winged insect" I can't even tell if it could be a bee or a fly.

I got it , it is a horned treehopper which is found in UK, US and India

From the wings, it must be an Auchenorrhyncha, likely a Membracidae species. So, I wonder how it could have bitten or stung anyone...
This one looks similar and occurs in India: https://www.projectnoah.org/spottings/16531012

I am also confused that , which type of eyes this treehopper has ..... These are not ommatidia not simple eyes but which type

It's a treehopper (membracidae) from the genus Oxyrachis. They do not sting or bite. Identifying the species is impossible due to the quality of the image, but here are a few:

O. tarandus

O. versicolor

O. sinensis


text:
Can a neuron produce electric or magnetic effect?
According to my view,As a neuron is an electrical impulse generating cell and through this impulse it carries information from brain to whole body or vice-versa, As here a very small electricity is generated so there must be some electrons which can produce an electrical or magnetic effect !

Any time charges move (current), electric and magnetic fields are created.
Neurons involve quite small voltages, on the order of 100 mV as well as small currents. Neurons process and transmit information through small changes in voltage, but they generally return to rest. That means, over a long time scale, the net electric and magnetic field is going to be zero: everything averages out to some steady state. Neurons and their fibers are made up of roughly spherical or cylindrical compartments, and voltage is going to be roughly constant within those compartments (because a solution with dissolved ions is a good conductor). That means the changes occur along the edges, and think of a sphere: if the same change occurs all around the entire surface, once you get sufficiently far away you're going to see all those fields as cancelling each other out. For a cylinder, the cancelling will occur in every direction except along the long axis of the cylinder.
That said, populations of neurons that have synchronous activity and are arranged in favorable orientations (like that of pyramidal cells in neocortex) can create detectable electrical and magnetic fields. Electroencephalography and magnetoencephalography are commonly used techniques to measure this brain activity (electrical and magnetic, respectively) from outside the skull.


text:
How much effort is it to establish a cytotox assay for cancer cell lines against a small number of possible compounds?
I am currently testing a series (5-10) of small molecule compounds against an enzyme that are intended as inhibitors. This enzyme is meaningful for cell proliferation.
Until now, nothing was active and therefore I could try to test the new molecules in a cytotox assay, to test if they are of any medicinal value. It is known, that these compounds have multiple modes of action and inactivity against the discussed enzyme does not necessarily mean that they are fully inactive.
I am wondering how complicated this is and if you're a beginner on this how long does it take to set up a growth inhibition or viability assay. Currently me gathered experience on protein expression and aspetic technique. But I have no experience with cell culture and sterile bench. My cell lines of interest are melanoma cell lines and or various brest cancer cell lines that are commercially available from multiple known suppliers. Do you think it would be possible to test this in a couple of weeks? I am willing to work 7 days per week if this will speed up the process. What is a realistic time frame to get first results? I would work with commercial inhibitors as control. I have no idea how time intense this process might be and would be interested about your insights.

Do not forget https://xkcd.com/882/ and https://xkcd.com/1217/

The answer to this is, of course, "it depends". There are quite a few factors at play here. As you don't seem to be experienced at cell culture yet, I strongly recommend that you find someone experienced to teach you. It is not something that is easy to pick up, as problems in cell culture (e.g. low-level contamination, cross-contamination, old cells, overgrowth, under-seeding) can be subtle and difficult to identify without experience.
It will take at least 4 weeks for experience to kick in, and cells aren't like bacteria in that they won't grow overnight to suitable levels for sub-culturing/experimentation; it'll take days to weeks of growth from a frozen vial to having enough to work with. This takes time alone. You'll need to be in some weekends anyway; when cells need to be split or fed, that's when you'll be in!
It takes time to learn the characteristics of each cell line, how fast it grows and how it behaves under certain conditions. If you get the conditions wrong, it will invalidate your experiment. Many cell lines will tolerate quite a bit of mishandling (e.g. growing to too high a density), but others will stop growing if you seed at too low a density and/or they become too confluent. This varies from cell line to cell line and even cell lines within a single type (e.g. melanoma) can be different from others.
You also need appropriate media for each cell line - many culture media use phenol-red as an indicator dye, but this is an estrogen analogue, which can affect growth of hormone-sensitive lines (often the case with breast cancer lines for example). Some lines are sensitive to glucose concentration or need a special supplement to grow. Knowing these sorts of things are critical to your experiments and is where experience kicks in. You will need to start by looking up the growth conditions of each cell line you plan to use.
Once you are experienced, it can be a fairly simple process. With enough cells and the right equipment (e.g. repeat dispense function multi-channel pipettes), I can easily set up 20 or more 96-well plates for viability assays (IC50 style) in a day. There are lots of other options on assays too, how long they take and what read-outs (MTT/XTT?, neutral red? cell count? luminescence or fluorescence-based) you use are variables too.

Thanks. I am now actually working of these experiments yet. For me, the biological part is not the challenge but the synthesis of Activa compounds. For the viability test I want to use the MTT assay. When it will show some.effect on the cells I want to proceed with apostosis tests and than metabolite studies by LC MS/MS. Keep thumbs pressed that my compounds will kill cancer cells.

Just wanted to follow up on this. I am handling 3 different cell lines at the moment. Cells are growing as expected, splitting freezing etc. everything seems to work. But I am not confident with my MTT assay results at the moment. The triplicates ain't looking good and I think it is because of the DMSO step to solubilize the furazan crystals.


text:
What are possible tweaks when a protein expression from a plasmid shipped on paper fails?
In my group we recently tried to express a protein necessary to perform an analytical method in BL21(DE3). The first two attempts did fail and no colonies did grow on the agarose. I hear, that the extraction of plasmid from paper and resoubilization is more prone to error in comparison to plasmid that is shipped in liquid form and this is a known issue. I am wondering what could be possible workarounds to try to still get it working. Until know, I have the following ideas:

Increase plasmid concentration
Decrease antibiotic concentration
Use a different strain like NEB 5 alpha

I am also wondering if the X-Ray treatment of customes service could facilitate in any way plasmid determination or is this an urban myth?

How did you prepare the plasmid from the paper? Did you do a transformation and a mini prep first?

No miniprep. I took the sheet of paper and cut it leaving a small portion as backup. Solubilzed the plasmid with 50 microliter TE buffer, resulting in a plasmid concentration of 11 ng/ml. Using 4 microliter plasmid with 25 microliter bacteria culture. Did not work. Than increased to 10 microliter plasmid, did not work. I observed now increased plasmid concentration of ca. 17 ng/ml after adding 100 DNSase free water destilled water and letting it stand over night.

Ok: My advice in these cases would be to first transform some standard competent cells (DH5a or likewise), cultivate them, make a prep and check your plasmid at least on a gel, ideally but restriction digest. Also make some glycerol stocks. So you make sure you have the right plasmid and you have a stock you can always go back to.

I will consider this after my current experiment batch. I will do 5 plates, 2 with Ecoli 21D3 and two with NEB5 alpha, two different antibiotic concentration and 1 plate just with LB media at the smaller antibiotic concentration. Therefore I should be able to reduce the amount of possible error sources based on the outcome of the experiment. When not a single of 4 plates show colonies I agree, the integrity of the plasmid is in doubt.

I think that your title here is entirely misleading - you aren't looking at protein expression in this case, simply transformation of the bacteria with a plasmid. Frist step is to test that your system works with your cells and plasmid and with known good cells and plasmid - beg/borrow known good competent cells and plasmid from a colleague and test them in parallel with your cells and plasmid. Note that you need competent cells for this, not just bacterial culture.

You're right. Goal is of course the expression after the trafo. Cells are competent. In the meantime I also got colonies and we will do a miniprep and check the plasmid.

In my case it helped to treat the filterpaper longer with miliq water to increase plasmid concentration. I also used more plasmid in the 3rd attempt and it worked. Miniprep was performed nevertheless. When a similar experiment is not working, I would recommend to increase plasmid concentration, increase incubation time and decrease antibiotics in that order. I tested the plasmid on agarose gel and further I sent the plasmid further to a sequencing service for validation. Seems all to checkout.


text:
For Cytotox studies of anticancer compounds: What cell line can I use to check if my compounds will also damage healthy cells?
I am about to perform cytotox studies on cells with my synthesized compounds.
I want to test my structures against MCF7 cells (breast cancer) as well as a melanoma cell line. MCF7 should be vunerable against my compound.
In the best case, the drug like compounds be especially effective against cancer cells but spare healthy cells.
I read that HeLa cells are used as a test subject, to see if chemicals are toxic to human tissue, especially when cosmetics etc. are tested. Could I use Hela cells therefore as a cell line to see if my compounds show off target effect against healthy, not overproliferating cells? Or what cells are used usually for this purpose and are they immortalized?

HeLa are highly abnormal and very very proliferative. Most cell lines are proliferative in some manner. This sounds like a question that would be best suited to your advisor as they should know the details of your project and what is available for you to use or what is normally done in your lab.

I am wondering what people are normally do, to explore off target effects? I am not the first person who is testing cm against cancer cells. I assume they just use cells from actually patients that are not cancerogenic?

There are a few options which depend on the stage of your project. You could try a line which isn't a breast cancer line (normal breast), you could try one missing receptors for your compounds (KO/KD or natural) , you could try another tissue type (e.g. ovarian), another cell type (fibroblast instead of epidermal), you could try normal human primary cells, cells derived from directly from biopsy. All are possibilities... Like I said earlier, it's a question for your supervisor, there is no "normally do" in this, it's something that is project specific.


text:
Volatile alternative to PBS that is compatible with LCMS analysis for cell lysis experiment
I have two chemical compounds that I put on cells, to see if they can diffuse into the cytosol. The compounds are similar in structure, but contain a key modification that alter the polarity completely. After the cells are incubated, they are washed with PBS several times to remove excess compound. PBS is used not to rupture the cells pre maturely. The goal here is just to see compound that actually went into the cell, not something that was there from the beginning. The final sample contains the lysate after protein precipitation with 2 parts methanol and 1 part PBS.
PBS however contains of Sodium, Potassium, Phosphate and Hydrogen phosphate and Chloride ions which are all non volatile and therefore may cause problems with LCMS detection or ion suppression in some cases. Therefore I am  wondering if it could replaced by ammonium formate at 154 mM concentration at pH 7.4? For washing and injection steps? (NH4Fa is also the buffer in the LCMS run) Could this be used as a substitute or cant PBS be replaced?