text:
Does sample buffer require EDTA for protein separation on SDS PAGE?
In sample buffer preparation we add EDTA, but if SDS-PAGE is for protein then is it necessary to add EDTA in sample buffer? What is role of EDTA in sample buffer for protein separation for SDS-PAGE.

In my opinion EDTA is dispensable. Common proteases like serine proteases do not depend on metal ions. As the post in this page suggests, there are EDTA free loading buffers. This discussion in researchgate suggests that EDTA is excluded from protease inhibitor cocktails because it may interfere in downstream steps. I am commenting because I am not able to verify the dispensability of EDTA, from any reliable source.

I've never used EDTA in sample buffer. The combination of SDS and DTT (or β-ME), along with the boiling step, is enough to completely denature and inactivate pretty much every enzyme, including proteases.

EDTA is a complexing agent for bivalent cations and is usually used to inhibit enzymes (proteases, nucleases) which need these ions in their active center to work. Without it they do not attack the sample.
The presence of EDTA in the sample buffer is interesting, since Laemmli's original paper from 1970 ("Cleavage of structural proteins during the assembly of the head of bacteriophage T4.") doesn't use it and I think it is not necessary.
The reason for this is simple: Protein samples are usually generated in buffers which contain different protease inhibitors, which protects the samples. Additionally the purpose of the sample buffer is to denaturate (at least for the standard SDS gels) the protein sample completely, so it will run uniformly. Both steps ensure that proteases are either inhibited or completely denaturated, which will both protect the sample.
The only reason I can think of for using EDTA is that it can help to destabilize your target protein, when a co-factor which is important for the structure is complexed by EDTA and thus keep it in solution after the denaturation.
There are of course ressources which state that EDTA is necessary for protein integrity, but they only say it is there to inhibit proteases by complexing metal ions (see here for example).
As a 2x sample buffer, I use the following recipe (this can also be made 5x if necessary), which contains EDTA, but can also be done without. If you prefer β-Mercaptoethanol, you can use it in the same concentration as the DTT (which has the advantage of being less smelly):
2x sample buffer:

4% SDS
20% glycerol
2 mM EDTA (ethylene diamine tetraacetic acid) - can be made without
100 mM Tris-Cl, pH 6.8
200 mM DTT (dithiothreitol)
0.1 % (w/v) bromphenol blue dye

AFAIK only metalloproteases use metal ions (Zn and Co); these are not the most common proteases that one would encounter.

There are some calcium dependent proteases. You may also want to inhibit kinases or phosphatases.

There are very few of Ca dependent proteases. I am not sure kinases need metals as cofactors (activation is a different thing; that's signaling). Alkaline phosphatase needs Zn. But I guess this is not as common as the dependence of nucleases on metal ions. In any case we do use a protein inhibitor cocktail while doing extraction. Loading shouldn't require EDTA; you are just denaturing.

@WYSIWYG I looked into Laemmli's paper from 1970, and he doesn't mention EDTA at all. My guess for the moment is, that it has been added later for unknown reasons. I think since the concentration is not high anyways (1mM final), there will be no harm if it is removed. I'll check that later and edit the answer.

Yeah I guess it is like one of those occult biolab practices of putting every coloured substance including copper sulphate solutions, covered in aluminium foil at 4⁰. Not sure though.

It's not unlikely. Although the colored dye here make sense as it makes loading easier.

I agree with the other comments but want to add something.
EDTA works as a stabiliser for bME. If you have bME in your 5x sample buffer and want to store it for a long time period add EDTA, otherwise you have a big loss of activity.
https://www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Sigma/Product_Information_Sheet/1/m7154pis.pdf


text:
How can I get brighter DNA bands?
I amplified my DNA with real time PCR. The amplification curve shows good amplification but not a single band is observed in my gel after gel electrophoresis. What would be the possible reasons?
I am attaching a photograph of my gel below. My ladder opened correctly but not a single band can be observed:

In which cycle did your sample come up? Are you sure it is your wanted product and not a primer dimer (which are visible on the gel)? What is you expected product size for your PCR product? How much DNA from your qPCR did you load on the gel?

almost at 30th cycle.yes these are primer dimers. i load about 5ul of pcr products in my gel.

my product size is 170 base pair

Do you know that you have product besides the primer dimers? How does the melting curve look like? There you should see two melting peaks - one for your product and one for the primer dimers. Besides that, 30 is relatively late, so there is not much template. You could of course precipite the whole PCR reaction and then concentrate it to see the product.

yes there are two melting peaks.but i quantify my pcr products with nanodrop method; quantification shows that signifacant amount of amplified dna is present

The top of the gel looks really red, are you using too much ethidium? Sometimes the background can obscure faint bands, and you can avoid that putting a small amount of ethidium in your DNA loading buffer instead of the entire gel.

will u please send me the recepie for gel preperation may be there will be a problem in my gel preperation method

i used 5ul of ethidium bromide

@tanzeela For which volume of gel?

@tanzeela Did you clean-up your product before measuring it on the Nanodrop? If not, you will measure residual dNTPs which will give you a too high reading.

Can you put a pic of amplification curve. Another thing. It would be good if you can resize your image so that they dont exceed 500×500px

@chris for 2% gel

@tanzeela I mean how much of your gel did you make - 50ml, 75ml....?

@chris for 50 ml

As you can see from the bottom of the gel, the brightness of the bands have reduced even for the DNA ladder/marker. The main reason for this is that when you are running an agarose gel containing EtBr, due to the positive charge of the EtBr, the stain runs to the negative chamber, against the direction of the DNA migration as @Chris has pointed out! Although some of the DNA will stain but the rest of the EtBr will migrate towards the top of the gel (where the DNA is loaded).
A quick remedy for this is once you ran your gel is to soak your gel in a separate container containing your stain and running buffer such as TAE. I do this for one hour and that makes a huge difference!! Also make sure you regularly change the buffer in your agarose gel running apparatus and don't use the same buffer for a long time since again as @Chris has correctly pointed out there is a reduced conductivity in the older buffers when they heat up so migration does not work as well! Obviously this response assumes that you have a good degree (amount) of DNA in your gel bands, which can be calculated by nanodrop once you extracted the DNA using an extraction kit, or whatever method you normally use. I hope this helps!
This answer is more suited to the title of the question since after reading the main question a bit more carefully I'm slightly confused! So do you see anything at all or your product is far too faint? could you tell us where on the gel, relative to the ladder you expect to see this band? are they the ones at the button of the gel, below the loading buffer?

Your assumption about the Ethidiumbromide is wrong. In solution it is charged positive so it runs to the kathode and against the running direction of the DNA. Some of the EtBr will intercalate into the double helix and stain it (although weaker the further the DNA migrates), the rest migrates as long as the power is on. This is why it is usually recommended to stain gels which run very long afterwards, as this increases the staining.

@Chris Ah, but I was told it is the pH effect that causes this since in my apparatus containing old running buffer this is worse than when made freshly and when I use the buffer in the positive chamber side of the apparatus and sock in EtBr, it doesn't stain. But I could be wrong! I can edit my response if thats best?

No, the pH has almost no effect on this. You will loose conductivity in older buffers as well when it heats up, so the migration is not working that good anymore. You could add EtBr also to the lower buffer reservoir, but this can get messy.

@Chris ah interesting! thanks! I shall edit my response!

source: Stack Exchange

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

sort: votes


text:
Is there an optimal composition and length for protein linkers in FRET?
I'm designing a protein that I'd like to use in FRET reporting.  General idea on the shape is:
FRETprotein1--Linker--CleavageSite--Linker--FRETprotein2.  I would like to know what AA are best for the linker, or does it matter?  I have heard SSG is a good choice, but when I started a lit review I become unconvinced.  If it doesn't matter, I may try and put more convenient restriction sites there.
Is there a good way to theoretically determine optimal length?  I realize that may change depending on which FRET protein pair is chosen.  The cleavage sites are between 15-18 AA long.  Unfortunately, I don't think I can give any more detail than that on the cleavage sites.
Response to answer:
So far there have been no FRET systems or tests published with the cleavage sites I want to study.  In fact, I haven't even heard of anyone else trying.  If I'm forced into a positive FRET probe, which I'm worried that I am, does anyone have any FRET pairings that don't interfere with Texas-Red on the spectrum?  My first choices were Venus and Cerulean, but if I have to make a positive probe then I don't think they are a good choice.

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


text:
High Current (Speed) Transfer Buffer Recipe
Does anyone know an effective buffer mix to use for high current Western transfers?  We are successfully using the vendor's premixed buffer to transfer a wide range of protein sizes to PVDF membranes at 1A/25V for 10 min.  We get great results with the vendor's expensive buffer.
I haven't been able to find a non-proprietary recipe that works.  The best ones we have hold the 25V, but then have a decrease in conductivity (increase in resistance) where they will start at 1A, but drop to 0.4A by the end of the 10 min.  The vendor's buffer seems to hold the high current and results in better transfers.
I don't know if it will be helpful to list all the variations I've tried in detail, but they have revolved around modifying a traditional Towbin buffer plus SDS, more glycine/Tris, or MgCl2.  These were all various suggestions from people around the department, but I haven't found much published evidence for a buffer under these conditions.
I realize 1A (and no I don't mean 1mA) is a lot of current, but the vendor's system works really well.  Any pointers on what I might try/add would be appreciated even if you don't have a worked out protocol.
Edit: Info from the MSDS indicates it has 3 reagents:
Listing of dangerous and non-hazardous components:
Proprietary Reagent K
10-20%
Proprietary Reagent EB II
5-10%
Proprietary Reagent S
1.0-2.5%
7732-18-5 water
50-100%
·
.....
Solvent content:
Organic solvents:
0.0 %
Water:
74.8 %
Solids content:
25.2 %

[I'm not sure if this MSDS info should be put here, just because I'm looking for someone who has already used a high current buffer they know the formulation for, not a guess to what I have.]

@leonardo I believe that even though Life Technologies (gotta keep up with the mergers) would call iBlot a "dry transfer," that it really seems like a semi-dry transfer with very nice pre-packaged cell set up.  I have used them, and the gels certainly seemed wetter than dry.  I feel like Licor as more of a 3-rd party non-player had an interesting analysis.

@leonardo although you get the same resolution/sharpness using precast gels and the standard Towbin's wet transfer method...

@MattDMo  I certainly have plenty of love for a classic Towbin transfer, but I'm in a high western throughput kind of place right now.  I'm doing about 4 gels and 16 blots a day through high-speed solutions (and getting good data), I was just hoping to find a way to bring costs down.

@AtlLED - I understand where you're coming from, I was just trying to clarify leonardo's comment about clarity. FWIW, I used to run at least that many westerns a day: I used Invitrogen precast gels, and did a wet transfer at 100V (constant) for 1hr@4C. One of the issues we found with the iBlot was its transfer efficiency of very small/large proteins - you can get one, or the other, but generally not both.

I would look both into the MSDS sheet and, if any, the patent. They should help you to identify the components but not the recipe.

Already did.  Obscured in propiatry.  Even gave huge ranges in what ever the mistory compounds were.

So after a lot of work in optimization, I thought I would post what worked best for me.  This buffer recipe was able to successfully transfer EGFR and insulin from the same lysate, and a clear band for both (large and small protein respectively). 10% SDS-PAGE gels were transferred at 1A for 10 min.
High Current Transfer Buffer

48 mM Tris
15 mM HEPPS
1.0 mM EDTA
1.3 mM NaHSO3
1.3 mM N,N-dimethylformamide
25 Mm gLY-GLY
20% Methanol (v/v)

I do hope this can help someone else, and want to site Garic et al as wonderful starting point.  Their publication was made after I asked the question, as @user4148 pointed out.

The SB buffer has been around for quite a while, and I definitely have used it successfully before for both agarose and PAGE gel buffers for some time. The only thing is that you have to run the gel in a cold room (high voltage equals high temperature) and that you have to optimize the timing and conditions so that your samples don't run off the gel.

I've used SB.  First, isn't it a running buffer, not a transfer buffer (or at least I've never heard of it being used in transfer), second I believe the formulation is still some what broad no?

If you look at the running buffer recipe it contains SDS, whereas the transfer buffer contains methanol. Using the same base buffer but adding the different reagent, I have used it before. However, you might have to try it with your specific recipe and running conditions to optimize it.

A recipe with SB buffer for western https://www.mcdb.ucla.edu/Research/Banerjee/protocols/Western-Blotting.pdf

My point wasn't that I couldn't find SB recipes, my point was I could find several.  I will give the linked one a try and see how it stands to optimization.  I still can't find examples of SB in high current low voltage situations (especially for protein transfer).  The one you linked notes 25V o/n, which is the opposite of what I'm trying for.

I am just pointing out that it can be used for transfer buffers as well. But interesting enough people seem to have mixed experiences with it. I think in general you probably don't want to run fast transfers if you are looking for very precise or accurate results because it is generally a trade-off that you need to consider.

Ah but there's the rub, I'm getting both precise and accurate results with the vendor's buffer.  I've compared it directly to an overnight slow tank transfer.

Probably because they have spent lots of time and money optimizing the product. Many of the proprietary recipes and protocols are not more or less complicated, just better optimized I guess...

Yeah I don't think a SB buffer is going to work.  I tried several formulations, and they all fared far worse.  Some even boiled.


text:
Cause of Crease in Drupe Fruit
What causes the crease through the pericarp of a drupe fruit (peach, nectarine, olive)?  It may just be in exocarp as @Ilan pointed out, but I assumed (perhaps wrongly), that the division continued through the pericarp.  I'm also not sure if this true generically for drupe fruit, so I'm not sure which phylogenetic step down to take.  Olives for example have a divot that seems to extend from the stem down, but it stops well before the midline of the fruit in most cases.
I assume the crease has something to the initial genesis of the fruit tissue, but I haven't found any arguments on it's formation.  A more technical term might be "cleavage," but that was after some searching and not coming up with more certain term.
Botany is not my strong suite, but I recently entered into a collaboration where we will be making some recombinant alpha viruses (which is within my strong suite) that will infect drupe tissue.  Finding good pathology papers has been difficult, and they tend to be specific (not trying to imply that's bad).  Can anyone point towards a reference text/article on the genius of drupe fruit if the cause of the "crease" is not clear?  Disruption of the crease seems to be a particular phenotype of infection for some strains, and I'm searching for the correct terminology and mechanisms that might be effected.
If I can get permission I will include photos of an infected vs control peach.  In the mean time to clarify what I mean, the line that seems to transverse from one poll to the other of the fruit:
Peach:

Nectarine:

On the sea mango it is pronounced, but on most other mango it seems like just a slight marking along the exocarp:

Even the coconut seems to have at least one:

What do you mean "crease" - a part of a normal state, or abnormal sign?

@Ilan Part of the normal state.  The outside of the fruit tends to have one line/partition/crease going down it from the outside.  I'm uncertain if there is a technical name, which is where I could be failing in my lit review.  I suppose I could put up a picture and circle what I mean.

I think you should change pericarp to exocarp (pericarp is all-out-of-stone term)

@Ilan I actually don't know if this just on the surface at the fruit or if it extends down to the stone.  That may vary based on species even.  I'm not sure on how many species will be tested, but distorting the line in peaches seems to be an unexpected phenotype.

The structure you are relating to is a seam (suture). This seam is formed during development of a carpel.
The indentation of the seam varies and depends on the climate and environment: for example, water stress will give a very deep suture -

Know any proteins related (screen via Western) or genes (q-PCR)?  We could grab samples in various stages during carpel development (though this particular set of trees might not make it to next year).

@AtlLED check this one out - http://www.ncbi.nlm.nih.gov/pubmed/19264761


text:
Does DNA being circular or linear directly affect the speed of DNA replication?
Let's say we have two DNA molecules of equal length, one belonging to a prokaryote and the other to an eukaryote. It's known that replication of the eukaryotic DNA is faster in this case. One clear reason for this is that linear DNA has multiple origins of replication whereas circular DNA only has one.
Now back to the real question: Does it matter for rate of replication whether the DNA is circular or linear? Does it contribute to eukaryotic DNA replication being faster in our specific case?
One thing I found in an old revision of this Wikipedia article is as such:

One reason that many organisms have evolved to having linear chromosomes is due to the size of their genome. Linear chromosomes make it easier for transcription and replication of large genomes. If an organism had a very large genome arranged in a circular chromosome, it would have the potential problems when unwinding due to torsional strain.

What I make out from this is that the rate of replication isn't directly affected. It's more closely related to avoiding other issues that arise from eukaryotic DNA molecules being typically longer than prokaryotic ones. But in our case where we declare both DNA to be of equal length, I'm assuming circular vs linear has no bearing on the rate of DNA replication.
So am I correct on my assumptions?
Edit in response to the answer below:

It's known that replication of the eukaryotic DNA is faster in this
case (where DNA molecules are of equal length) because eukaryotes
have linear chromosomes whereas prokaryotes have circular ones

If it makes sense to form a sentence like this, presenting linearity as the cause, then it's enough to satisfy my definition of directly affected in this case. For example, it's easy to make this claim if you present the number of origins of replication as the cause. I looked at the textbook you mentioned and the quantity of origins of replication is in fact mentioned this way.
Conclusion:
I went ahead and did a bit more reading in that textbook. What
I've understood is: An eukaryotic linear chromosome has multiple
replication origins rather than a single one in order to compensate
for its much larger size. So while the shape might be a factor
(possibly, not certainly), the primary and "direct"
reason is the difference in size, not the shape.

The current Wikipedia entry has modified this to "In an organism with a very large genome, circular chromosomes could potentially cause problems relating to torsional strain.[citation needed]". The much more cautious language presumably reflects the lack of citation in support of the postulated effect of torsional strain. Naively I would imagine less torsional strain in a larger circular chromosome, but in any case, there is no evidence presented that torsional strain is actually relevant.

@David Yeah, I agree that it's definitely not a highly credible claim the way it currently stands. But still it doesn't directly matter for the real question since it was only a (somewhat baseless) idea for why circularity/linearity might not have a direct impact on the speed of replication.

I'll see if I can find anything about this when I have a moment. As you may be aware, many closed circular DNA genomes are replicated by the rolling circle method — which in itself may have different characteristics to the standard replication method.

@David Thanks. I guess if we consider the method of replication as well that might further complicate reaching an answer. It looks like this question does require generalizing and neglecting a bit to reach any answer at all.

I am not sure if I well understand what you mean by directly affected.
I will list some possibilities below (For reference you can see any genetics textbook, I use, Genetics: A conceptual approach by Pierce, but I guess any textbook would do).

Porkaryotic polymerases (usually processing circular DNA) have a higher nucleotdie per second speed than eukaryotic polymerases (processing linear DNA). Does this mean that shape directly affects speed? In this case I would say no. It's just a difference between prokaryotic and eukariotic polymerases. Some prokaryotes have linear chromosomes and their polymerase will still be faster than eukaryotic polymerases.

One prokaryotic chromosome has only one origin of replication, while one eukaryotic chromosome has several of them, so that eukaryotes parallelize even in a single chromosome. Does this mean that shape directly affects speed? In this case, I would say yes. Due to steric effects, the shape of the chromosome has to do with the ability of having several origins of replications, even if the lengths are the same. Edit: After Ved's comment, I realized that this is also not necessarily true. Not all circular chromosomes have one single origin of replication; archaea have circular chromosomes with more than one origin of replication

Linear chromosomes are easier to unwind. Does this mean that shape directly affects speed? I would say, yes for large chromosome and no for small chromosomes. Edit: After David's comment, I realized that I am not really sure that linear chromosomes are easier to unwind. A Wikipedia entry states this, but I gave a look at the paper they referenced and found no real statement about this.

Importantly, if the DNA molecules you compare are one prokaryotic and one eukaryotic and they are very short, I would bet my 5 bucks on prokaryotic (circular) being faster, because in absence of all the parallelization the eukaryotic DNA polymerases are slower than prokaryotic ones.

Could you provide a reference or detailed mechanical argument in support of your assertion that linear chromosomes are easier to unwind. This would seem to be the crux of the matter. What you say and what you would bet is not the point. The Wikipedia entry acknowledges lack of support for this sort of assertion. Please provide it if you want your answer to be considered seriously.

Actually, after your comment I realized that no, I do not have a reference. I checked tha paper referenced by Wikipedia, but actually I wasn't able to find a statement supporting my assertion (which originally I took from OP question, who was also referencing wikipedia). It is possible that I lost some important passage in the paper, but at the moment I would say it doesn't support the assertion. Thank you for pointing this out, I edited my answer accordingly.

Well in that case, could you provide a reference for steric effects affecting the quantity of replication origins? I looked at two textbooks but couldn't locate anything relevant.

While I was looking for a reference to support the steric effects affecting quantity of replication origins, I actually found out that archaea have circular chromosomes and more than one origin of replication. I edited my answer to incorporate this.

While the question may not have gotten a definite answer (might be impossible anyway), I think the findings here are indication enough to reach some conclusion at least: It's not possible to generalize and say that the shape of DNA will certainly have an effect on the rate of replication. Number of replication origins on the other hand seems to almost certainly always have a direct effect on the rate. I will mark this as the answer, thanks.

I agree that it may be impossible to generalize. But I want to point you to this paper I just found: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6056714/ . Give a look at the "separation mtDNA" section. It deals with the structural problems with the circular mtDNA, and also references some work on prokaryotes. Maybe it will not answer your question, but it may be useful for information.

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


text:
Gene of interest number of copies in transformed K. pastoris
When transforming a K. pastoris strain, is there a way to predict how many copies of the gene will be integrated following the homologous recombination? More specifically is it something that a molecular biologist is able to control in a simple way, or, is it usually left to the chance (and determined afterward using gene sequencing)?
Alternatively : if I want to transform an electrocompetent wild type strain such as Pichia X33 using a plasmid vector, is the multiple integration an iterative process I control or is it random?)

Edit: Thank you all for the answers. Finally if I understand well the process is pretty random but you can control it to some extent. @gaspanic: I keep the idea of using fluorescent reporters and FACS for that ;)

Do you mean if you target the same locus in a diploid/polyploid species or do you mean off-target integration?

Yes if I am targetting a same locus in haploÏd or diploÏd specie.

I only worked with with S. Cerevisiae and as long as you use 50 nt flanking regions it should only go to your region of interest unless your background has defective DNA repair mechanisms etc. In my lab we typically don't do whole genome sequencing just to verify a transformation. With conventional transformation techniques you only target 1 site in a diploid because the efficiency is so low, but with the cas9 enzymes you could integrate in both loci of a diploid.

Also your schematic won't work because the background will already have zeocin resistance.

Out of curiosity, is there a specific reason for not verifying systematically the transformation of final clone by sequencing? Apart from cost and time reasons? It is not the first time I see that. I guess yes, in the future CRIPR would be more used. (You are right about my schematic, it was only for illustration purposes)

I should mention we always verify the correct integration with PCR and gel electrophoresis. It also depends on what you're doing. For simple gene deletions in my lab we didn't do it. That's because the transformation efficiency is so low and using 50 nt homology flanks is so precise for targeting that the correct integration was likely the only recombination event. Also with yeast you can easily and rapidly test many independent transformations so rare events wont contribute much. But if you have a mutant with modified heterologous gene expression, whole genome sequencing would be important.

Thank you very much for all these precisions

To predict the number integrations is probably not possible, but you can favor multiple integrations, if that's what you are after. At least in Saccharomyces cerevisiae integration of a first plasmid appears to stimulate the insertion of additional plasmids at the same site (1). It is therefore possible to favor multiple integrations (2) by increasing the DNA-to-cell ratio in the transformation, thereby increasing the chance that any competent cell receives more than one copy of the plasmid, which could then integrate in tandem. The easiest way is to simply use more DNA, but depending on your cell competence you might need to compensate this increase by using less cells for your transformation (or simply plate less cells on each plate) as you will probably also increase the overall colony yield.
One way to actively select for multiple integrations is to use a competitive inhibitor for the selection marker. Again, in Saccharomyces cerevisiae transformations with HIS3 selection can be combined with 3-AT (3). This allows you to inhibit HIS3 activity just enough to only permit the growth of cells that have integrated a desired number of copies of the plasmid. Of course, this method is far from fool-proof, as you might also select for mutants that have increased HIS3 expression for other reasons. Edit: I actually found one paper doing this in K. pastoris (4).
Finally, if your implementation allows it you can facilitate the screening for multiple integrations by including a fluorescent protein-expressing cassette in your plasmid. That way you can easily screen for different numbers of integrations by fluorescence microscopy or FACS.
Sources:

Orr-Weaver et al. (1983): https://mcb.asm.org/content/3/4/747
Plessis and Dujon (1993): https://doi.org/10.1016/0378-1119(93)90172-Y
Wikipedia (retrieved 15 Aug 2020): https://en.wikipedia.org/wiki/3-Amino-1,2,4-triazole
Menéndez et al. (2018): https://doi.org/10.2144/00295rr05

Welcome to Biology.SE! How would using less cells compensate for having more DNA — did you mean more cells? This looks like a good answer, but answers are much more likely to receive a favorable response if you [edit] to include supporting references (primary literature is best). Without that support, your answer is indistinguishable from opinion. This 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. Thanks!

Thanks for your comments! I clarified the text a bit and added links to relevant papers. If you still think I should clarify it better, please let me know!

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


text:
Problem with an analogy (Cell Membrane)
I'm studying biology for the first time seriously. I've been reading OpenStax College's material.
In the section of "The Cell Membrane", I have the following problem:
Choose the answer that best completes the following analogy: Diffusion is to ________ as endocytosis is to ________.

filtration; phagocytosis
osmosis; pinocytosis
solutes; fluid
gradient; chemical energy

I chose number 1, but the site says the correct answer is number 2. Could someone please explain why?

[EDIT]
I chose number 1, because diffusion and filtration were passive ways to transport, while endocytosis and phagocytosis are active. I didn't not take in to account that 2. fitted better, because the osmosis part was confusing me.

I'm not sure why this question has 5 upvotes. This is a homework question with no attempt at working out an answer.

@James, sorry I thought of writing something else, but since this was an analogy I didn't know what else to put. Is it necessary to delete this?

Why was the answer to this question moved to the comments?

@AnaGalois It sounds like your question is actually about osmosis rather than anything else. If you are confused about osmosis, then ask a question specifically about whatever is confusing you about osmosis. This sort of MCQ format isn't a good for SE. See this meta discussion.

Osmosis is the diffusion of water.  Pinocytosis is the endocytosis of liquid/water.

While your answer is undoubtedly the one the question is looking for, osmosis is not the simple diffusion of water.


text:
Cell organels during cell reproduction
When one studies cell division, usually the process is explained through what happens with the DNA of the cell, particularly, that in eukaryotes the nucleus dissolves. But what happens to the rest of the organelles? I read here what happens to the ER and Golgi complex, in particular I'm wondering what happens to the mitochondria, in which part does it reproduces? do they make a parallel process? or do they get dissolved?
$%---------
%I started wondering about this because I was reading Lynn Margulis' article _Bacterial Bedfellows_, and it baffles me that in such little time an organism can just coexist with an attacker and turn it in a _good relationship_, how ever it's not clear to me how cell reproduction might work in those cases.$

During mitosis the cellular organelles that have their own genetic information reproduce independently. Other organelles like the Golgi complex disintegrates and then form all over again in the daughter cells. Other organelles that are smaller and more numerous like ribosomes are divided between the two cells. Meiosis is the same thing only that there is crossing over in the nucleus of the cell.

So mitochondria reproduces at the same time as the rest of the cell?

Mitochondria can reproduce on their own. I'm not sure how this is controlled exactly, so maybe it happens at a different time point. Also note that DNA replication (or reproduction if you will) does not happen at the same time as cell replication (mitosis/meiosis)

@Nicolai 'Also note that DNA replication (or reproduction if you will) does not happen at the same time as cell replication (mitosis/meiosis)'... In cell cycle, S-phase and M-phase are two distinct phases where the cell divides and DNA replicates receptively. Also the cell cycle almost in all standard text based on the pattern of division is termed Mitosis/Meiosis (not the M-phase). A slight wording issue.

@Nicolai It is controlled with AMPK https://biology.stackexchange.com/a/1094/54743

authors: Валерий Заподовников


text:
Prokaryote or just a small worm
In the following photos can be seen a thread-like (almost like longkou vermicelli) things. At first I was convinced that it was cyanobacteria, however a classmate said it look more like a little worm, and other person said it look like it was probably nothing (nor prokaryote nor worm). I can't find a way to settle what it is, how can I know?

Zoom 40x
This is from another sample

EDIT:
Could the first one be lyngbya? I found this website and the second to last picture kind of looks like what it is in my picture:
And also could my second picture depict nostoc bacteria?

The first picture shows something different than the two following picture, doesn't it? Did it move? What is the zoom? You might want to review a little bit your knowledge about the diversity of eukaryotes (incl. unicellular eukaryotes).

If it doesnt have a mouth and anus and organs it's not a worm.

@Remi.b I added that the zoom is 40x. It didn't move, at least not enough for being noticeable. I agree that I need to study about the diversity of single celled organisms, but why do you say eukaryotes? is that a hint? I thought it had to be a procaryote since it sort of look like a very long bacillus. Also, I was convinced it was cyanobacteria, but now I doubt it

What was the source of these creatures?

@Tytoalba Pond water... very dirty pond water

@AnaGalois Because you talk about worms! A worm is a eukaryote. You uploaded twice the same picture. The first picture was maybe the most interesting you might want to make sure to upload it as well.

@Remi.b Right!!! I'm sorry. Yes  I'll upload it again, however it is from a different sample

This would be a suggestion rather than a precise answer.
Your sample being pond water a highly probable candidate of the first two photographs would be Euglena sp., though the red eyespot is not visible.
They are freshwater Excavates   and according to Washington State Lake Protection Association a common pollution-tolerant genus in Washington water bodies.

Another picture reference: A website

I think Euglena is a good guess for one of the things in the picture, but I don't think that's what OP meant by "like longkou vermicelli"

:P I missed that.

Thank you. I also added another picture I found on the web, perhaps those two are related?


text:
What's the proper way to determine whether dose response curves +/- another variable (besides dose) are statistically different?
I have an experiment wherein I measured uptake of a certain molecule into cells when delivered using Carrier A vs Carrier B. In other words, for example, I delivered 1, 2, 3, 4, 8, and 10 nmoles of DrugX to 6 wells of cells per dosage, HOWEVER in 3 wells DrugX was delivered using Carrier A and in the other 3 wells DrugX was delivered using Carrier B. I would like to statistically determine whether the dose-response curve I get when I used Carrier A for delivery is any different than the dose-response curve I get when I used Carrier B for delivery.
My initial thought was to run a paired T-test, but then I realized I would have to report results at each point and I would like to report on the curve as a whole. My next thought was to use a two-factor ANOVA, however from what it looks like, when it determines whether or not there is variance due to the change in carrier it looks at the average and variance of all the responses. Another thought I had would be to calculate area under the curve, but I'm not sure how. I finally settled on running a nonlinear regression on the Carrier A data, then on the Carrier B data, then as the dataset as a whole and compared the resulting fit curves. I got a nice p-value, however the $R^2$ values are lower than I'd like (about 0.9) because the curves didn't model especially well.
What would you do? What is the industry standard?

For the ANOVA you would want to focus on the interaction terms, those would tell you if there is a difference in the effect of drug depending on carrier for each level of the drug. Same thing for a regression, you should be comparing models that allow for interactions with carrier, not just a main effect of carrier (it's possible you did this, I can't really tell from your brief description). If the drug effect follows some expected pattern, a generalized linear model might be better than a nonlinear regression. I can't speak to industry standards, however.

Might be worth asking this on stats.stackexchange, too? I have seen paired t-tests with results at each point reported in the literature, but I dislike this quite a bit. I suspect the answer depends on what aspect of the dose-response curve you care about: is it the saturation value? The 50% peak response?

What is the expected shape of the dose-response curve in this case? Or if there is no expected shape, what do the curves look like?
If there's a transformation of the data that makes it look linear, I would also suggest fitting a generalized linear model (i.e., with a non-linear link function) and looking for a significant main and/or interaction effect of Carrier B, as Bryan Krause suggested. If there isn't such a transformation, there are likelihood-ratio tests you can do using non-parametric regression, but you might not have enough data (see e.g. EDGE from the Storey lab).

Good call on not using multiple t-tests. The reason being, assuming a 95% confidence interval, each time you run the test you are "allowing" a 5% possibility of making a Type I error - this means that you incorrectly reject the null hypothesis in favour of the alternative hypothesis. If you run one t-test, there's a five percent chance of making an error. If you run two t-tests, there's a ten percent chance of making an error.
Between a regression model, or ANOVA, it does not matter. As the mathematics behind each are pretty much equivalent.
What you really want to look for, is the slopes of both models. If the two models have a statistically significant difference between their slopes, then this is indicating an interaction effect.
Two examples of (some, but not all) possible interaction effects:

Carrier A & B have a similar effect on dose-response at 2mmol. But as the dosage approaches 10mmol, Carrier B causes a dose-response that is nearly double that of Carrier A.
Above 5mmol, Carrier B creates a greater dose-response. Below 5mmol Carrier A creates a greater dose response. At 5mmol the dose-response is nearly equivalent.

I agree, t-tests are not appropriate. I don't think an ANOVA is great here either because you are not concerned with an average. With dose response curves, you typically are more concerned with an effective dose or a lethal dose derived from a model fit to the data.
There are well-developed standards for this type of analysis as it is common in pharmaceutical development, ecology, and enzyme kinetics. The most commonly used and acceptable models used to analyze dose response data are 3 or 4 parameter logistic models that end up looking s-shaped. You can find a brief overview of this here.
If you are familiar with R, there is an entire statistical package developed to compare dose response curves called drc. This is a general description of the package and its use. This paper describes more concisely the use of the package and the theory involved.


text:
Molds and fungi usually found on book pages: identifying and preventing further spread
I know I can use bleach to kill most of the spores that live on my walls. I know I have to keep humidity below a certain level and temperature above another to prevent the spread of mold. I have read tons of websites on how to treat 'wall infestations'.
It's the mold that affects my books that worries me the most.
Especially now that I've discovered a very enticing market for used books. You can find bargains but sometimes the books smell "of mold".
I would like to know more about the kind of molds (and/or fungi or other bio-monsters) that usually feast on paper. The paper that makes the pages of a book.
Can someone point me to any kind of resource with answers to the following questions?

What are the most common biological causes of page deterioration?
1b. Can they be identified without lab instrumentation?
Can they spread from one book to another, and if yes how and how fast?
Can they be neutralized in some way (stopping the spread), or should one just burn the moldy books to avoid contagion?

I seek knowledge to protect knowledge.

Mold can grow in Extreme environments: they can live in very low water activity level, sometimes as lower as 0.71, live in refrigerator temperatures, these characteristics make it difficult to kill them thoroughly.
You can search "paper deterioration" in Google or library, then you can find lots article about this.
1) Causes
Foxing and other white, fluffy coating or light yellow powder on paper caused by: Eurotium rubrum, E. repens, Aspergillus versicolor, Penicillium solitum, and Penicillium decumbens. (Karbowska-Berent, Joanna, Joanna Jarmiłko, and Jolanta Czuczko.
Source: "Fungi in Fox Spots of a Drawing by Leon Wyczółkowski." Restaurator. International Journal for the Preservation of Library and Archival Material 35.2 (2014): 159-179.)
Black spots: Monilia, Aspergillus, Penicillium, Mucor, Stemphylium, Hormodendrum, Fusarium, Chaetomium, Byssochlamys
Source: Beckwith, Theodore Day, Wilbur Harold Swanson, and Thomas Marion Iiams. "Deterioration of paper: the cause and effect of foxing." Publ. Univ. Calif., biol Sci. 1.13 (1940): 299-356.
2) Spreading
For their spread, I think the mold can spread everywhere, but i did not find any resource about it.
3) Method that kill the mold
Drying the affected paper objects and applying 70% ethanol are the most preferred options to stop active fungal growth.
Source: Sequeira, Sílvia O., Eurico J. Cabrita, and Maria F. Macedo. "Fungal Biodeterioration of Paper: How are Paper and Book Conservators Dealing with it? An International Survey." Restaurator. International Journal for the Preservation of Library and Archival Material 35.2 (2014): 181-199.


text:
How do PER/CRY complexes accumulate in the cytoplasm?
The Period/Cryptochrome (PER/CRY) complex is a heterodimer that transcriptionally regulates circadian rhythm. When it is phosphorylated, it is transported to the nucleus and it inhibits a transcription factor that regulates the synthesis of its own subunits (negative feedback).
Research on the circadian molecular clock speaks of an accumulation of PER/CRY complex in the cytoplasm during the day phase (see review The circadian clock: pacemaker and tumour suppressor, Fu and Lee, 2003). Why does this complex not move to nucleus during the day? In other words how is the phosphorylation-dephosphorylation switch controlled?

Welcome to Biology SE! In order for people to grasp your question, please clarify the question by explaining the acronyms (i.e., get rid of them :) and work a bit on the grammar as it is very hard to read. a citation may help a big deal as well.

With the edit, the answer is there in the question itself.

@WYSIWYG Maybe my edit wasn't clear--PER/CRY builds up in the cytoplasm during the day because it is not phosphorylated (and therefore not transported to the nucleus). At night, it is phosphorylated and negatively regulates itself. I know the answer has something to do with blue light and Cryptochrome but I don't actually know the reason

@Luigi added an edit to mention the phosphorylation


text:
What are the main principles of the Bacteriological Code?
Although the Bacteriological Code is an important guideline for taxonomists and microbiologists, describing the criteria for delineating species and higher taxa of the prokaryotes, I, and plenty of other microbiologists with me, would not be able to describe the main rules by heart. I am also not able to find a clear summary anywhere on the web, and reading and understanding the Code itself is currently taking quite some time.
Furthermore, does the Code say anything about taxa mainly or solely composed of uncultured strains based on 16S sequences, apart from the addition of Candidatus to currently uncultured single species? I am specifically referring to higher taxa, large groups of sequences.


text:
how to reconcile PDB structures' sequences via Uniprot references?
I’m trying to reconcile structures  in PDB entries with their  sequences as reported in their chemList.polymer.dbref entries.
E.g.: considering structures for HIV integrase:
PDB:3OS0 references UNP:P14350, focusing on integrase (len=391)
>sp|P14350|752-1143
CNTKKPNLDAELDQLLQGHYIKGYPKQYTYFLEDGKVKVSRPEGVKIIPPQSDRQKIVLQ…NRTVSIDNLKPTSHQNGTTNDTATMDHLEKNE
PDB:2B4J references UNP:P12497, focusing on integrase (len=287)
>sp|P12497|1148-1435
FLDGIDKAQEEHEKYHSNWRAMASDFNLPPVVAKEIVASCDKCQLKGEAMHGQVDCSPGI…LLWKGEGAVVIQDNSDIKVVPRRKAKIIRDYGKQMAGDDCVASRQDED
Why should these sequences be so different?!
and in case you think I'm focused on a bit of noise: there are 42 other HIV integrase structures all referencing UNP:P14350, and 43 other structures referencing UNP:12497 (and 79 others referencing UNP:Q76353, very similar to UNP:12497), 22 referencing an entirely different UNP:Q72498.

None of them is integrase:
P12497
Protein    Gag-Pol polyprotein
Gene       gag-pol
Organism   Human immunodeficiency virus type 1 group M subtype B (isolate NY5) (HIV-1)
P14350
Protein    Pro-Pol polyprotein
Gene       pol
Organism   Human spumaretrovirus (SFVcpz(hu)) (Human foamy virus)
Even if you meant the integrase product of the polyprotein, the second one is not even HIV.
From Wikipedia:

Human Foamy Virus (HFV) is a retrovirus and specifically belongs to
  the genera Spumavirus. The spumaviruses are complex and significantly
  different from the other six genera of retroviruses in several ways.
  The foamy viruses derive their name from the characteristic ‘foamy’
  appearance of the cytopathic effect (CPE) induced in the cells.[1]

good catch, @wysiwyg!  first, yes, i have been focusing on just the integrase bit of the polyprotein (eg, that's what the 1148-1435 BLAST qualifier does).  and  i've purposefly been including PFV in my analysis because "... PFV is very similar to HIV integrase and provides an excellent model for studies of retroviral integration".  but obviously i need to check organisms before aligning their sequences!  thanks again for your answer.


text:
Do animals have different taste preferences within the same species?
Humans (especially children...) seem to dislike certain foods or drinks, that other humans seem to like. Common examples are coffee, french cheeses, olives, milk, fish and cabbage. Are there examples of animals either wild or in captivity that show the same behaviour? For example a group of primates that eat something except one or two members of the group? And if so, any clues to the evolutionary aspects of such preferences?

Certainly.  Just for an example, one of my dogs likes carrots and apples, the other won't touch them.

In captivity this seems to be observed quite often - almost every pet holder (at least the ones that live with at least two pets of the same species) will confirm this. In my cats, for example, this also is not restricted to taste but also includes smell. Generalising this to wild living animals and an evolutionary perspective is really interesting. Good question!

Yes. Intra-species variation in food preferences has been observed in

Rabbits (Bilko et al. 1994)
Copepods (Rieber 1982; Berghe and Bergmans 1981)
C. elegans (Abada et al. 2009)
Earthworms (Bonkowski 2000)
Butterflies (Jermy et al. 1968)
Beetles (Sarker et al. 2013)

I am sure most pet owners will find the existence of this variation quite intuitive. No too cats have the same food preferences.
Why such variation exist?
I can think of several possible (non-exclusive and non-exhaustive) reasons for why such variation exist?

Individuals compete. If preferences vary, it allow specialisation (inspired from @John's comment)
Phenotypic variance (whether genetic, environmental or other) is not necessarily maintained by some kind of balancing selection. You may observe purely neutral variation in preferences.
Genetic variance can explain variance in food needs and therefore it is good that preference can be plastic
Same as above but for environmental reason. Say, an individual may have different need based on its environment.
One must learn to recognize good food. There might be a tradition among family of liking specific types of food.

Thanks for the links, fascinating (+1). Do you have any ideas about the evolutionary aspects?

if your taste varies a little bit from your main competitor (others of your species) you are likely to find food they have overlooked, several variations on taste is more evolutionarily stable than just one species wide preference.

@RHA see edit in my answer

Thanks, as your answer is the best of all given, I am inclined to accept it ;-)


text:
Is Black mulberry always deciduous?
I have a black mulberry (Morus sp.) in my garden (or 2 actually). According to the information I could find it is a deciduous tree. The trees in my garden do indeed lose their leaves. However, as most native trees start to color yellow/brown/red in september and have lost all leaves by now, the mulberry doesn't. It remains green until the frost kicks in and only then drops its (still green) leaves. It almost looks like it doesn't want to drop its leaves, but is forced to so so. This can be seen at the picture below. Hence my question:
Is mulberry also deciduous in warm winters such as the mediterranean? And is it normal that leaves are still green when dropped?

The trees are generally deciduous in temperate regions, but trees grown in tropical regions can be evergreen.
According to wiki:
https://en.wikipedia.org/wiki/Morus_alba

Thanks, I thought as much but couldn't find a reference.

WARNING: This answer is very tangential to the question. Confused Morus sp. with Rubus frutticosa (don't use "common name in questions"! luckily I wrote the answer using scientific names). So, as addendum: some trees tend to lose some leaves later (in my region, such trees can recover easily with broken branches).
"Always" is "always" a strong word in biology.
You describe your mulberry (Rubus sp.) as deciduous: the leaves are changed every year.
Many plants (especially trees) lose leaves in autumn/winter and they get new one in late winter/spring, few plants are marcescent, e.g. various oaks (Quercus sp.), so the leaves dry in winter and the plant lose it just when the new leaves appeaers. Some plants lose the leaves in summer.
Mulberries, like various "small plants", tend to keep the leaves until they form the new one (eventually, replacing all the shot). In my zone mulberries tend to keep old leaves until the new one are already developing.
Why this habit? Mulberries lives at borders of forests, so they try to have more energy/sun when the tallest tree lose leaves. On the other hand, being shrub and tending to radicate easily, it keeps safer in case of frosts, and it is also not a long living organism (as "plant", not as clone).
Note some decidouos young trees have also the same habits: they keep their few leaves in winter, to try to catch up. Being young also means few investment for the original plant, so it could be still a gaining habit. Look this winter on undergrowth.
Not so different to strawberries and many grasses, which replace the leaves in springs. But they are not "decidouos", just because they doesn't have a trunk or just they replace the entire shot the shots.

I agree one should always use both names (added it to the question!). Still, I am not sure where your confusion comes from. If I say Mulberry and link to the wikipedia page on Morus sp., why do you think I am talking about Rubus sp.? Which BTW are called blackberry or raspberry, but never to my knowledge mulberry (do you have a source for this name?). Finally, Rubus sp. are always (small) shrubs, never trees. And finally: your entire answer lacks references.


text:
When do the spindle fibers attach to the chromosome
At what phase does this occur in mitosis (or even meiosis); some text books say prophase while others say metaphase.

Would you like quoting which book said what?

Happens in prometaphase.

Your confusion comes from the inconsistent terminology in the literature. The attachement of the spindles to the chromosomes happens in the Prometaphase [1, 2]. The name already suggests that this is not one of the classical phases is cell division, but lies somewhere between Pro- and metaphase. Wikipedia states:

Prometaphase is sometimes simply included as part of the end of prophase and early metaphase.

If you really need to stick to the classical scheme of pro-, meta-, ana- and telophase, I would personally suggest to treat the attachement (not the formation!) of spindles as a part of the metaphase.


text:
Growing E. coli at room temperature?
If I were to do a blue/white selection of transformed E. coli on LB agar ampicillin plates at room temperature (23⁰C) for about 2 days and 18 hours, will I run into the issue of satellite colonies or any other issues?

Why do you do it this way and not use the classical overnight incubation followed by 1-2 hours in the fridge? Works usually pretty fine.

I normally incubate overnight at 37 C, but I was just curious if it's possible to achieve the same results at room temperature over the span of a weekend (when I am unable to come in to my lab and refrigerate the plates).

I have never tried this for blue/white selection, I have overexpressed protein a 18°C to have a higher yield of soluble proteins. I think it will not do harm, as the bacteria grow much slower. If you don't see a clear selection, you can still put them into the fridge.

ampicillin degradation is a real issue at room temperature for more than 1 day. YOu will see a ton of satellite colonies and false negatives

Yes, once I did it (pBR322's fault) and I did see some satellite colonies.

Since b/w screening is a means to  selection, you want maximum selection pressure. This will be obtained by incorporating all optimal conditions. The many enzymes of the lac operon are going to function best at their optimal temp.
Why would you do this? If it is because you can't go in the next day it's better to do it when you're available. You're only going to doubt your results when you get them.

Blue/White screening is not a selection step (unless you count picking the right colonies as selection).

@chris you are correct it's just a means to selection

You will definitely get satellite colonies. Ampicillin breaks down after about a day, especially exposed to light. Kan, keeps pretty well at room temp, but that does you no good :)


text:
How to compute the regression of individual fitness on individual phenotype
Consider a population structured in groups of two individuals. Individuals' interactions follow an additive prisoner's dilemma:
\begin{array}{c |c |c|}
& C & D \\
\hline
\text{Cooperate (} C \text{)} & b -c & -c \\
\hline
\text{Defect (} D\text{)} & b & 0 \\
\hline
\end{array}
where $b$ is the benefit and $c$ is the cost. The payoffs are to the player on the left. I need to calculate  the regression of individual fitness on individual phenotype, $\beta (w_i, p_i)$, where $p_i = 0$ if $i$ defects and $p_i = 1$ if $i$ cooperates. Note that I need to calculate the regression within groups rather than between groups.
I thought that the right way of calculating $\beta(w_i,p_i)$ would be to vary the strategy of the focal individual while keeping the strategy of the other individual constant (my thought was that the other individual becomes the environment for the focal individual). Because switching to cooperation causes an individual to lose $-c$ in fitness (if we keep the strategy of the other individual constant), I thought that $\beta(w_i,p_i) = -c$. But, to my surprise, $\beta(w_i,p_i) = -b -c$. To show why, McElreath & Boyd ("Mathematical Models of Social Evolution", p. 242) draw the following graph:

Let $V(X|Y)$ be the payoff of individual $X$ when it interacts with $Y$ (in other words, $X$ is the focal individual; $Y$ is the other individual). I understand that McElreath and Boyd calculated the regression coefficient by computing $V(C|D) - V(D|C)$ --- as opposed to $V(C|C) - V(D|C)$, where you keep the strategy of the other individual constant. My question is why this is the right way of calculating $\beta(w_i,p_i)$.

I totally forgot to say what $V$ was. I fixed this now. Thanks for pointing it out.

If I understood your second question correctly, I want to calculate the regression of fitness on phenotype. The example only deals with two phenotypes, cooperate and defect; and the population only contain two individuals.

@Remi.b : I rewrote the question and added a graph explaining the example. Hope it's clearer now.

I am not sure I understood the question. Let me know if this helps.
Case: N=2, freq=0.5
Let's assume that the frequency of those who cooperate is 0.5. The slope of the regression line (which R.squared is equal to 1 as we have as many data point than degrees of freedom) is by definition $\frac{\Delta w}{\Delta p}$. You defined $\Delta p = 1$. What is $\Delta w$ then?
Well, given that one cooperate and the other defect (as indicated in the fact that the frequency of cooperators is 0.5), then one will have the payoff of a cooperator that is facing a defector and the other one will have the payoff of a defector facing a cooperator. The cooperator has a fitness of $w_0 + b$ and the defector has a fitness of $w_0 - c$. I think you may misread the payoff matrix. To know the fitness of someone with strategy $i$ when facing someone with strategy $j$, you look that payoff at row $i$ and column $j$.
Therefore, $\Delta w = (w_0 - c) - (w_0 + b) = - (b+c) = -b - c$, and so the slope is  $\frac{\Delta w}{\Delta p}=\frac{-b-c}{1} = - b -c$
Case: N>2, freq=0.5
In such case the slope of the regression will necessarily be lower than $-b-c$ because some cooperators will meet other cooperators, increasing their fitness (=highering the points on the right hand side of your graph) and some defectors will meet other defectors, decreasing their fitness (=lowering the points on the left hand side of your graph)
At equilibrium
The only stable equilibrium is when the frequency of cooperators is 0. In such case, there is no variance and we cannot even talk about regression.
Conclusion
The slope is not necessarily $-b-c$. It is $-b-c$ for the very special case where we have two individuals, one cooperate, the other one defects. In any other scenario the slope $s$ of the regression is lower. More specifically, the slope $s$ of the regression for any $N$ and for any $freq$ is $0 =< s < -( b+c)$

In a case in which the frequency of cooperators should be 0.5, I can see why the regression coefficient has to be $-b -c$. My question arose because McElreath and Boyd derive the regression coefficient without making any assumptions of the frequency of cooperators in a group. If we assume that we can have any frequency of cooperators in a group (0, 0.5, and 1), it seems that the regression coefficient can have value $-c$.

I agree with you that $-b-c$ is not the slope of the regression in any case. Yes, in some cases the slope of the regression can be -c. I did not calculate the slope of the regression for any $N$ and frequency but it probably is not very hard I guess. See edit.

Here is my mistake, I think.  $w_i$ can be written in terms of $p_i$ in the following way (note that $p_i$ can only assume two values, 0 and 1):
$w_i (0) = w_0 + kb$
$w_i (1) = w_0 -c + (k-1)b$
where $w_0$ is the baseline fitness and $k$ is the number of altruists in the group (so, $k$ can be 0, 1, or 2). When we are calculating $\beta (w_i, p_i)$ the question we are trying to answer is "for a given group in the population, what is the regression coefficient $\beta (w_i, p_i)$?". This implies that $k$ must be constant because, otherwise, we would be considering individuals from different groups. Accordingly,
$\beta(w_i, p_i) = w_i(1) - w_i(0)$
$\beta (w_i, p_i) = -b -c$
For $\beta(w_i, p_i)$ to be equal to $-c$ we would need to consider the payoff of cooperators from different groups.

Following up on the equation Remi.b suggested in a comment below, we can write the regression coefficient to a population structured in groups of $n$ individuals. More specifically,
$w_i(0) = w_0 + [k/(n-1)] b$
$w_i (1) = w_0 -c + [(k-1)/(n-1)] b$
With this definitions, we get
$\beta (w_i, p_i) = w_i(1) - w_1 (0)$
$\beta (w_i, p_i) = -c - b/(n-1)$

$b$ and $c$ doesn't have the standard definition as given by Hamilton. The fitness of an individual who cooperates is usually computed as $w_c = w_0 + x PO_{cc} + (1-x)PO_{cd}$, where $PO_{cd}$ is the payoff of an individual who cooperates when meeting an individual who defects and $x$ is the frequency of cooperators in the population.

My impression is that, in the example above, your equation yields $w_c = w_0 + x (b-c) + (1-x)(-c) = w_0 + xb - c$. The chance of a cooperator meeting another cooperator will be $(k-1)/(2-1)=(k-1)$, which gives the equation I wrote. Am I missing something?


text:
Why is squeezing not recommended after a needle prick?
I've read some interesting things about the risk of infection after a needle prick (30% - hepatitis B, 1.8% - hepatitis C and 0.3% - HIV), but I've also saw that the local squeezing is not recommended.
It's true that I've also found some articles that considered that there is no difference if you squeeze or not, but most of the articles I've read (in my native language) specified "Do not squeeze!!!". By squeezing I mean an act of extracting the blood by pushing the skin near the needle-prick.
Why?

If you've found relevant literature, don't just mention, cite!


text:
DNA barcoding and real-time PCR
I recently read an article on how DNA barcoding was used to identify species present in health products.
I also read an article about how Real-Time PCR was used to identify meat species in meat products.
I'm curious as to why one method is used over another. I know that DNA barcoding uses mitochondrial DNA to detect species, and Real-Time PCR uses species-specific primers to detect species.
Am I correct in thinking that real-time PCR cannot identify unknown species, but can tell you if the DNA matches a specific species you're looking for? For example, if I test for E. zebra but find something else, I cannot identify that unknown species and more testing is required?
Whereas in DNA barcoding, all I do is match the results against a huge list of known species in a database, which means I can identify unknown species without providing any prior knowledge?

PCR cannot identify species - this is usually done by sequencing. The only way to do so (for known species only) is to use species specific primers which either give you amplification of DNA or not. So can make a relatively simple yes or no decision on the presence of the species, but you cannot go further beyond this.

You've done a pretty decent job of answering your own question, but there are a few things that can be elaborated on.
The general convention is to use the mitochondrial Cytochrome C Oxidase 1 (COI) gene for barcoding animals and chloroplast genes (rbcL, matK, and trnH-psbA) for plants. The three biggest reasons for selecting these genes are 1) their ubiquity; 2) the presence of many copies per cell, so it's easier to PCR amplify the sequences; and 3) mitochondrial and plastid DNA has a much higher mutation rate than nuclear DNA, making it possible to uniquely barcode very closely related organisms. This method is incredibly sensitive, reliable, and easy. The original DNA barcoding publication can be found at doi: 10.1098/rspb.2002.2218
Thanks to the efforts of consortia like CBOL, BOLD and iBOL, there are now many millions of reference sequences databased, covering pretty much any multicellular organism you are ever likely to encounter (assuming you don't work on a deep-seq sub or something).
As you say in your post, there are significant limitations to using qPCR to identify species. You have to know beforehand what it is you are looking for, and you are left with a simple yes-no. This is not to say there is no place at all for qPCR in the species identification game, however. The method is very fast, and in cases where you are only interested in detecting a single species or small number of species, it's much more convenient than getting the DNA sequenced (for example, during quality control in a meat processing plant, doi: 10.1080/02652030701584041).

DNA Barcoding is a method or a protocol which, as you have already mentioned, uses genetic markers to identify species. The barcodes can be "scanned" using PCR.
Now Real-Time PCR is just a quantitative version of PCR. You don't need that to look for qualitative aspects. However in a mixed sample you may be able to calculate percentage constitution of the sample by each species, using Real-Time PCR.


text:
What plant is this, seen at the Allan Gardens?
I tried a couple of apps on my phone, but I could not identify this flower and plant species.  These are at the
Allen Gardens Conservatory, at the beginning of Spring in Toronto.

This is a Schizanthus "orchid" (actually not a real orchid) called Schizanthus pinnatus ("butterfly orchid" or "poor man's orchid").

It is native to Chile but the plants are also cultivated as ornamentals. It is commonly known as the "small butterfly" ("mariposita") or "small, white butterfly" ("mariposita blanca").

Description

Annual plant of 20-50 cm in height
Stem: Glandulous-pubescent
Leaves: 2.5-3 cm long; pinnatisect (divided into 6-8 pairs in oblong-linear segments, entire or separated).
Flowers: white, pink or violet, 2-3 cm diameter, arranged in panniculate inflorescences, sometimes dichotomous.


text:
What is the Attributable Risk of birth defects in Zika infected pregnancies?
Vital Signs: Update on Zika Virus–Associated Birth Defects and Evaluation of All U.S. Infants with Congenital Zika Virus Exposure — U.S. Zika Pregnancy Registry, 2016
A recent CDC report shows that 10% of laboratory confirmed Zika pregnancies in the US had "birth defect reported."  However, I cannot seem to find an attempt to compare this to a base rate in the US of a similar population.  Is there existing information or a way to determine the Attributable Risk (difference in rate of a condition between an exposed population and an unexposed population).
10% sounds scary, but without a base rate, it's not very meaningful in the context of determining how dangerous the Zika virus is.
EDIT (for clarification):
I'm primarily concerned with the method in which birth defects were collected. 'Clinical experts reviewed reported information to ensure that each fetus or infant with birth defects met the criteria of the USZPR case definition.', but this seems to have been done only with the Zika-positive groups and not with a Zika-negative group. They compare it to a pre-Zika baseline, in which inclusion to the surveillance was different and determination/clinical review of each case was different. Did they (or another research group) perform clinical evaluation of birth defects during the surveillance period on Zika-nagative pregnancies using the same methodology?

The article cited in the question (Vital Signs: Update on Zika Virus–Associated Birth Defects and Evaluation of All U.S. Infants with Congenital Zika Virus Exposure — U.S. Zika Pregnancy Registry, 2016) gives the base rate for the definition of "birth defect" that they use:

The baseline prevalence of defects consistent with those that have been observed with congenital Zika virus infection was approximately 2.9 per 1,000 live births in the pre-Zika years (4). The initial findings from the USZPR represent an approximate twentyfold increase in Zika virus–associated birth defects among pregnant women with laboratory evidence of possible recent Zika virus infection, with an approximate thirtyfold increase in brain abnormalities and/or microcephaly.

The reference for that background is Baseline Prevalence of Birth Defects Associated with Congenital Zika Virus Infection - Massachusetts, North Carolina, and Atlanta, Georgia, 2013-2014.:

These data, from Massachusetts (2013), North Carolina (2013), and Atlanta, Georgia (2013–2014), included 747 infants and fetuses with one or more of the birth defects meeting the case definition (pre-Zika prevalence = 2.86 per 1,000 live births). Brain abnormalities or microcephaly were the most frequently recorded (1.50 per 1,000), followed by neural tube defects and other early brain malformations† (0.88), eye abnormalities without mention of a brain abnormality (0.31), and other consequences of central nervous system (CNS) dysfunction without mention of brain or eye abnormalities (0.17). During January 15–September 22, 2016, the U.S. Zika Pregnancy Registry (USZPR) reported 26 infants and fetuses with these same defects among 442 completed pregnancies (58.8 per 1,000) born to mothers with laboratory evidence of possible Zika virus infection during pregnancy (2). Although the ascertainment methods differed, this finding was approximately 20 times higher than the proportion of one or more of the same birth defects among pregnancies during the pre-Zika years.

According to Estimating the Number of Pregnant Women Infected With Zika Virus and Expected Infants With Microcephaly Following the Zika Virus Outbreak in Puerto Rico, 2016. the background of microcephaly in Puerto Rico was around 12 cases per year, while they estimated 180 cases would occur in the presence of Zika.

We estimated an IQR of 5900 to 10 300 pregnant women (median, 7800) might be infected during the initial ZIKV outbreak in Puerto Rico. Of these, an IQR of 100 to 270 infants (median, 180) may be born with microcephaly due to congenital ZIKV infection from mid-2016 to mid-2017. In the absence of a ZIKV outbreak, an IQR of 9 to 16 cases (median, 12) of congenital microcephaly are expected in Puerto Rico per year.

This is modeling, not pure observation, but was based on observed risk. The paper is online full-text (I think) so you can follow through the assumptions if you want.

thank you for the information.  Unfortunately, the answer I'm looking for is slightly different.  I just want the rate of "birth defects" in the US in a way similarly defined by the CDC.  The rate of microcephaly alone is far to narrow when considering all "birth defects", and I would be hesitant to use Puerto Rico rates, because of obvious differences when compared to the continental US.

That answer seems to be in the first paragraph of the article you cited, so I'm not sure exactly what you are looking for. I updated my answer anyway

I'm primarily concerned with the method in which birth defects were collected. 'Clinical experts reviewed reported information to ensure that each fetus or infant with birth defects met the criteria of the USZPR case definition.' but this was done only with the Zika-positive groups and not with a Zika-negative group.  They compare it to a pre-Zika baseline, in which inclusion to the surveillance was different and determination/clinical review of each case was different.  Did they not perform similar tests during the surveillance period on Zika-nagative pregnancies?

With the moving goalposts and the changing demands, it seems like you have some weird axe to grind, and comments are not the place for it, so you're on your own.

your information is helpful.  Typically, attributable risk is done by tracking the exposed and unexposed population in the same way.  I'm interested if this has been done by the CDC or some other group.


text:
How do we feel temperature?
Recently I watched a Youtube Video by
Veritasium
He said we are actually feeling the rate of heat transfer, not the temperature.
But how do we feel temperature?
How is feeling temperature different from feeling heat and heat transfer?

Welcome to the Biology Stack Exchange. Please take the [tour] and carefully read through the [help] to learn more about the site, including what is on-topic and what is not, and how to ask a good question.

@Mesentery - although your links are highly useful, new users are likely overwhelmed and do not know what you are after. If you have certain critiques, please make them explicit, especially to new users. Posting a standard welcome message with all sort of links within the site may have its merits, but is arguably not the best thing to do. Other users have started doing that and have eventually been discouraged. Specific pointers are, imho, better.

@AliceD I have seen that this user has not got the informed badge. I just want him to guide him about how this site works. So, if he/she asks further any questions, they are also on-topic on the site.

@Mesentery - that is awesome. Mentioning this explicitly will help this new user even more I think. Thanks for all your good work here and for your efforts. I'm not aiming to criticize, just trying to be constructive. Cheers.

After all that being said, a summary would be - did you try finding an answer? 'Provide details. Share your research', else it is likely to be closed as a low effort question. I'm voting as one.

use google.  "nerve temperature response" = https://en.wikipedia.org/wiki/Thermoreceptor afaik they have different ranges for different precision/speed of response. the topic takes 30 40 minutes of going on good pages to be knowledgeable on for the rest of your life, and one day you can use all your knowledges as a combined power of generalist science knowledge. research is good.


text:
Can 'human' become a genus due to space colonization?
I have read that during the Second World War, some mosquitoes got trapped in the London underground railway system. The mosquitoes never got out and eventually they became a new species by themselves.
I had a similar thought. In the next few centuries, if humans could, in theory, colonize other planets like Mars, Proxima Centauri and beyond, then the environments there are not the same as Earth. So, in the long term, humans who would be born and who would grow up on Mars, for example may become more and more suited to Martian conditions than Earths.
Now, when early humans explored and ventured into new geographical areas, they did change characteristics, but we are still one species Sapiens. But living extraterrestrial, is a whole new thing. The gravity alters, the entire atmospheric composition does. So that is going to have some significant changes on humans.
So, my question is: is it possible that in millions or even billions of years, if humans expand to space, there may arise separate species of humans? And would this new emergence of human species actually result in humans moving one step up the taxonomical ladder: becoming a genus?
EDIT: To avoid confusion and create speculations at the answers, I should specify that I am talking about a very particular case: if Sapiens are living in different planets, then is there a chance that Sapiens will become a new genus, and that Homo can be taken one step higher in the taxonomical order? There would still be Sapiens on Earth, but considering the environmental changes that could happen here too, humans then can be drastically different from humans now. So the question is: can 'Sapiens' become a genus?
Thanks to @tyersome and @jamesqf for pointing this out.

Welcome to Biology.SE! Thank you for taking the [tour], but please also go through the help pages starting with [Ask] questions effectively on this site and [edit] your question accordingly. In particular, questions must be clear — how are you defining "human"? I would say the genus Homo is equivalent to "human" in which case your question is trivial. In addition, as written your question is speculative and thus will generate opinions rather than factually based answers — this makes the question off-topic as currently written. Thanks!

Human, or in Latin, "Homo", already IS a genus.  It's just that the other species in the genus -  Homo habilis, Homo erectus, Homo neanderthalensis, &c - have died out.

@tyersome thanks for the note, I edited it.

Define "species". IIRC there's been research that has shown that there's a bigger genetic difference between white and black people than there is between dogs and wolves, and there's a gigantic amount of physical variation between different breeds of dogs, who are all considered one species.

@nick012000 I don't know about the difference compared to wolves/dogs, but there is significantly more diversity within each human "race" than there is between "races", which is to say that any two black people or any two white people are generally more different than the average of all white people are from the average of all black people (and the same is true of all "races"). All that is to say that there is not much genetic difference between black people and white people

@KevinWells My point is that the definition of "species" is basically arbitrary and there's no concrete way to tell whether or not two distinct populations are different species or not.

@nick012000 does not a common definition of species require that mating between Earthling and Martian could not produce fertile offspring?

@MatijaNalis Dogs, wolves, and coyotes are all considered different species and they can produce fertile offspring.

@nick012000 Dogs and wolves are not considered different subspecies, dogs are just a domesticated sub-species. You are right about coyotes though, they are considered a separate species, but can interbreed with wolves. It's possible that they should therefore be considered sub-species, but it really just shows how complicated species distinction is

The concept you are referring to is speciation and it has been well studied in a wide variety of different natural organisms. I suppose here we are talking about the biological species concept.
The overall answer is yes it is possible, but critically depends on a few different factors. The reality of speciation in the wild is very complex, but these are some things to consider:
Genetic isolation
If two groups, such as your Martian colony and humans on earth isolate from one another, then for speciation to occur, there needs to be a significant level of genetic differentiation between them. This means substantial differences in the kinds of genetic variants found at positions along the genome.
Genetic differences are eroded by post-isolation gene flow – in your example, that might mean a spaceship flying back to Earth with Martian colonists, who then have offspring with people on Earth.
Although there is plenty of evidence that speciation with gene flow can occur, the general rule of thumb is that increased gene flow means a longer divergence time is required to fully speciate.
In nature, this is often caused by some kind of geographic barrier to gene flow, such as mountains or rivers forming, but it can also be caused by morphological differences, such as variation in sexual appendages. Of course, in our example, this barrier to gene-flow would be the large and difficult to traverse distance between the Earth and Mars.
Divergence time
You alluded to some kind of separation time, and you are right to be talking on the scale of millions of years. Speciation can occur extremely quickly; in Lake Tanganyika cichlids, it has occurred probably within the last 15,000 years. Humans have created whole new species of crops, such as maize, within the past 10,000 years. There is even evidence of some fish speciating in 3000 generations.
However speciation is often a much longer process. For example humans and chimps were thought to speciate in 4.5 millions years.
Of course, there is an interaction with several other factors. All other things being equal, less post-isolation gene flows results in a shorter time to speciation and vice versa. Stronger selective pressure between the different environments leads to a more rapid accumulation of genetic differences.
As Konrad Rudolph correctly points out, divergence time is strongly related to generation time, with all other things being equal, a shorter generation time, results in faster speciation.
Selection pressures & differing environments
I think the last main factor governing speciation is how different the Martian colonists environment was from that on Earth.
Different environments can lead to natural selection occurring in opposing directions in the two populations, leading to ecological speciation. Speciation can proceed without differing environments, where neutral drift in allele frequencies can eventually cause speciation, but this will be a long process.
Speciation will occur much more rapidly if there is a start difference in environment and selection pressures between the two groups.
So in conclusion, given enough time, genetic isolation and differential selection pressures (or some combination of the above), it is plausible that a new species of human could form. However, given the time-scales required, it seems a bit unlikely to me.
EDIT
I think it is worth pointing out what @Jaquez said in the comments. If current terrestrial humans split from an extraterrestrial source to form another species, it would be named as another species within the Homo, such as Homo extraterrestrialis, for example. The addition of a new species does not move the group Homo up to become an e.g. Tribe or Family.

Your answer currently implies that speciation time is the same for all species. This is very crucially not true, it depends on effective population size and generation length. The former is small in humans and the latter is very long, both of which imply that splitting into separate species will take much longer for humans than it takes for cichlids or maize, independent of selective pressure.

Which part of the answer do you think implies that? It wasn’t my intention.

Yeah, I didn’t think it was intentional. But the paragraph under “Divergence time” reads as if the numbers from cichlids and maize could be used as a proxy for humans (or rather, they could if it weren’t for “natural” vs artificial environments — which anyway only applies for the maize example, not for cichlids). That said, the 3000 generations you cite sound about right as a (very rough) ballpark estimate.

Right, I understand. In hindsight that was quite a tangential point and potentially confusing, thanks for noticing it.

Excellent answer, +1.  However, I feel it would benefit from clarifying that speciation would not, as the OP asked, move either Homo or sapiens up the taxonomical ladder.  It would result in a new species within Homo, for example Homo extraterrestrialis.

I do not believe it will happen.  There are multiple roadblocks:
First, speciation time is measured in generations, not years.  The human generation time is long, the 3000 generations mentioned in another answer for a fish translates to nearly 100,000 years.  Are human populations not going to interbreed over a time span that long?
Second, the speed of speciation is a function of the evolutionary pressure.  We radically alter our environment and change the whole driving force--genes which make one more or less able to survive the rigors of the environment have almost no effect anymore.  To the extent that human evolution will occur it will be mostly towards things that favor reproductive success--things which attract mates, a desire for children, contraceptive unreliability.  These things aren't going to be substantially different on Mars than on Earth.
Finally, we are already to the point that we can make designer babies to some degree, although we can't yet do it with the sort of reliability needed to ethically do it on humans.  I find it inconceivable that we can remain a technological species (as would be required to have a population on Mars) without reaching the point that we can freely rewrite the genetic code at least during reproduction.  If there is a speciation event it will come from the lab, not evolution.  It's possible that some day there will be a modification developed that is a substantial upgrade but which is not compatible with existing humans--those with the modification can only breed with others that have it.

Indeed. For such a speciation to occur, we had to first terraform Mars to a degree where a non-technological human population can survive, and then we would need a total societal collapse on both planets, so that space travel would become impossible for a long long time. But even then, in those 100000 years humans would either rediscover technology and space travel, or die out.

"is it possible that in millions or even billions of years" - so definitely.  Chimps and Humans diverged 5-7 million years ago.  As for @vsz, that's not true.  You would just need to have limited interaction.  If you are imagining societies with regular flights between the planets then maybe not.  But even on Earth we get 'ring species' that can interbreed at the boundaries while populations at the ends cannot.

@JasonGoemaat A requirement for speciation is that you have populations that do not interbreed.  That means a lack of mobility.  I'm questioning how you could have a population on Mars that for a vast period of time doesn't interbreed with the population of Earth.  If they fall too far for spaceflight they also fall too far for survival in the Martian environment.  And that doesn't even address my other points.

As others have noted, the key concept is speciation. Imagine some H. sapiens living on Mars. How are they going to breathe? Presumably we have a nuclear reactor generating power, which in turn is used to make oxygen. What is it breaks down? Those who can tolerate low oxygen levels might do better.
Here on Earth we invest a lot of energy in building bones and things to resist gravity. On Mars there may be less need to do this, and more need to be efficient with oxygen and tolerate higher levels of CO2. So some mutations that would be sub-optimal on Earth might be beneficial to the colonists.
Couple this with isolation, as others have mentioned, and the two populations may drift apart, leading to the eventual emergence of H martiansis (or whatever.).

A driving force producing natural selection is not required for speciation.  Separate two identical populations and give them identical environments and they will eventually diverge due to random changes in the genome.  One of the main factors causing this is in the proteins in the coating surrounding the egg.  They are like a lock that sperm have the key to.  The selective pressures preventing change are simply that a male cannot reproduce if it cannot find a female with compatible proteins. Small changes aren't a problem, but the drift over time would lead to speciation with no other changes


text:
What are the definition of pecking order?
Pecking order is the phenomenon that dominating individual usually eat first. It is commonly observed in animals. In animals, it is often due to the fact that the strong individual won the fights for eating first, or is able to win those fights in future.
This is what I used to learn: "a pecking order is about access to resources decided before the access as the result of a competition, not just straight competition". For example, the order of food-feeding in bird is sometimes decided by brightness of feathers, without any direct competition (fight) for the food.
However, many other sources equate pecking order with dominance hierarchy, or the metabolic evolutionary pressure, namingly some individuals have an evolutionary advantage over others, making them more fit to survive .
For example, bacteria can has dominance hierarchy (a.k.a pecking order). Supporting literature include this.
See also history of "pecking order", pecking order of plants, and another article on dominance hierarchy.
I am not sure if my understanding is correct and if this viewpoint is universally accepted -- is there a consensus on the definition of pecking order?

I think you have it wrong - a pecking order is about access to resources decided before the access as the result of a competition (usually strength, but not always, brighter feathers, bigger antennae can work for this), not just straight competition (being able to move faster). Voting to close for homework.

@bob1 How does this look like a homework? I get your point. I used to have the same viewpoint as yours but later I found that "pecking order" is equivalent to "dominance hierarchy" according to Wikipedia. I don't know who to trust. If you have one or two reliable reference I'll be appreciated!

It looks like homework because of these sections of the homework part in the help: "A question that addresses a basic biology concept that may seem trivial to biology professionals" and "Include details of how you have attempted to form a solution. We will not do your homework assignments for you but we will point you in the right direction if you have made the effort to research the topic first."

@bob1 "Include details of how you have attempted to form a solution" makes a question good, not bad (or close-worthy). All questions are encouraged to include the attempts to solve the question.

But you don't show how you have attempted a solution - you've found a paper on bacterial competition, but you don't provide why you think this is appropriate to support your postulate, or any other evidence that you actually know what a pecking order is. Also your "pecking order" (1st) link is literally on peck strength in birds - nothing to do with pecking order at all. These together mean this is a low-effort homework post.

@bob1 I don't get your statement. First you said this question "looks like homework because the question include details of how I have attempted to form a solution"; then you said I did not attempt anything.

@bob1 Anyways, I must restate that this is clearly not a homework question. So it seems like that you have a different definition for "pecking order". May I again kindly ask your reference or sources? Most sources I found uses a similar definition to Wikipedia

No, what I said was that you need to provide details of how you have attempted to find a solution - read the link in my second comment. You haven't provided details. A pecking order is an informal hierarchical social dominance order where individuals higher in the order have more dominance or authority over those lower in the order. An example might be a family - Parents -> elder child -> younger child. A basic ecology or biology text will have a similar definition. This means again, homework as based on the "trivial for expert" part I quoted from the help page.

@bob1 You said that my first link is irrelevant to "pecking order". Could you please again provide reliable reference that Ebbe 1913 paper is irrelevant to pecking order?

My apologies I mis-read and looked at the citing paper, my Norwegian being somewhat lacking. However, I believe that the publication should be this one from 1921: Beiträge zur Biologie und Sozial- und Individualpsychologie bei Gallus domesticus (Contributions to the biology and social and individual psychology in Gallus domesticus) . Hønsenes stemme. Bidrag til hønsenes psykologi translates to "The voice of the chickens. Contribution to the psychology of chickens", which doesn't quite look right. However, I haven't been able to view the originals.

"Earliest" is a bit problematic; all species on Earth have been evolving for the exact same amount of time. In terms of generations, complex eukaryotes with long life spans have been evolving for a shorter time than many single-celled organisms.

@BryanKrause For example, the "earliest" language is created by Sumerian, according to many of the main stream anthropologists. In this question, the word “earliest” means the same thing. Maybe for biologists this word is a bit ambiguous. Of course, you are very welcome to suggest an alternative word or expression.

@HighGPA Well unless you expect to find this from the fossil record, the issue is that "earlier" things like Sumerian don't exist any more to be observed.

@BryanKrause Probably I should change the question to "approximately when did organisms evolved the property of pecking order?" Sometimes scientists and theoretically deduce an approximate time even before the actual evidence is discovered. Does this sounds better?