text:
Predicting the optimal sample size for a nested experiment
I need to test the effects of different treatments on plants and I would like to know the optimal sample size (number of plants that I need to sample). The variable I need to measure is the average number of insects present in a single plant. In the experiment, I have 3 main treatments and, in each treatment, I have 2 sub-treatments. The information I can provide are the following:

Average number of larvae per plant: ranging between 0.03 and 0.1
Standard deviation: ranging between 0.09 and 0.28
The desired $Z$-score is 1.96
I have no idea about the minimum detectable effect size but I can guess it could be 10%

Looking online, I have found different formulas that return different results. The first formula was:
$$n_1=\frac{(\sigma_1^2+\sigma_2^2/K)(z_{1-\alpha/2}+z_{1-\beta})^2}{\Delta^2},$$
where:

$\Delta =|\mu_2-\mu_1|$ is the absolute difference between two means,
$\sigma_1$, $\sigma_2$ are variances of mean #1 and #2,
$n_1$, $n_2$ are sample sizes for group #1 and #2,
$\alpha$ is the probability of type I error (usually 0.05),
$\beta$ is the probability of type II error (usually 0.2),
$z$ is the critical $Z$ value for a given $\alpha$ or $\beta$, and
$k$ is the ratio of sample size for group #2 to group #1.

With this formula, comparing these two means (0.03 and 0.09) and using a standard deviation of 0.09, I am getting $n = 35$ for each treatment.
A different formula I found is:
$$n=\frac{2\sigma^2 Z^2}{d^2},$$
where

$n$ is the required sample size per sub-treatment group,
$\sigma^2$ is the estimated variance,
$Z$ is the $Z$ score, and
$d$ is the minimum detectable effect size.

With this formula, using a mean of 0.03 and a standard deviation of 0.09, I am getting $n > 6000$ for each treatment.
Could you suggest me a simple and efficient formula for calculating the sample size?

Power analyses depend on 4 numbers: alpha (false-positive threshold for significance testing), power (probability to reject the null hypothesis given all the other parameters), effect size (how big is the difference between groups, relative to the variability), and sample size.
Given 3 of these numbers, you can calculate the 4th.
If you want to calculate the sample size, you need to know what effect size you are designing your experiment to detect. This isn't optional or esoteric, it's absolutely critical to the power analysis. The effect size most often used for "t-tests" and similar is the ratio of the difference in means to the standard deviation. There are alternative forms if you expect the standard deviation to be different between groups.
You also need to choose your alpha and desired power. You haven't mentioned either of these in your post. There are some typical conventions, like alpha = 0.05, and power is usually at a minimum 0.80, though many would argue it should be higher, at least 0.90.
In your first calculation, you've set the difference in means to 0.06 and the standard deviation to be 0.09, so your effect size is 0.06/0.09 = about 0.67. This is often considered a moderate to large effect size.
In your second calculation, I don't know what exactly you've done, because you haven't written out any of your numbers, but I'm guessing this is where you've used your "10%" number. 10% of what, though? An effect size is not a percentage, it's a ratio. It can't be "10%". But if you've set it to 0.1, or 0.006, or something like that, this is a much smaller effect, and your power analysis should tell you that you need a much larger sample to detect this effect. Seems like that's what you got.
I'd recommend using a tool like G*Power to calculate sample size, but right now you are very lost and not understanding what you are undertaking, so I would really recommend to step back and sit down to read extensively about what power analysis is and means. This is not just a formula to plug in, this is critical to designing your experiment. Open up a statistics text book and don't move forward until you understand.
My preferred way to do power analysis is with simulation, because simulation lets you see what a power analysis is really doing and lets you understand more about your data. To simulate a power analysis, you generate synthetic data under the assumptions you have, like means and standard deviation for your different groups, then you actually do a statistical test on these data. Use your desired alpha threshold, run many many independent simulations (1000 to 100000 times), and note how often your p-value is smaller than this threshold. That's your power. You can change the sample size and see how the resulting power changes.
While running simulations, check whether the assumptions you've made about your data make sense! For example, if your mean is 0.03 and your standard deviation is 0.09, how often do you see negative numbers in your simulation? How often do you see non-integers in your simulation? Do negative numbers or non-integers make sense for a count of insects? No? Aha! Then you are violating some assumptions of the tests you're doing, and you probably shouldn't be doing t-tests with means and standard deviations, but rather using a statistical model that deals with count data.
If this all seems too complicated, it's time to consult with a statistician.


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Biggest change in size from birth
Which mammal has the largest change in size from birth to fully grown percentage wise?

I  guess it maybe a marsupial for example red kangaroo.

If the answer below addressed your problem, please consider accepting it by clicking on the check mark/tick to the left of the answer, turning it green. This marks the question as resolved to your satisfaction, and awards reputation both to you and the person who answered. If you have >= 15 reputation points, you may also upvote the answer if you wish. There is no obligation to do either.

@JM97 is correct.
According to National Geographic >>
The gestation period of a Red Kangaroo is 33 days and it's baby at birth weighs a mere 0.03 oz. To put it into a more precise perspective, on birth, the baby weighs 1/100,000 of an adult red kangaroo's weight.
Second place goes to the baby of a Giant Panda; at birth it weighs 3.2oz which is 1/900 of an adult giant panda's weight.

How about platypus?

Although platyplus babies are almost the size of a jellybean, their baby to adult ratio isnt much. Because adult platypus usually extend upto 15 inches.

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text:
What does it mean when observed genotype frequency is different from expected genotype frequency in Hardy-Weinberg equilibrium?
As I understand it, if a population is at Hardy-Weinberg equilibrium, then the genotype frequencies should be $$p^2 + 2pq + q^2 = 1,$$ given the allele frequencies of $p$ and $q$, which you can figure out if given the number of homozygous recessive individuals in a population (such as 1/10,000 babies are born with PKU, so $q = \sqrt{1/10000} = 0.01$).
What I don't understand is what does it mean if your expected genotype frequencies do not match your expected frequencies. For example, here is a textbook problem where there are two alleles, $V$ and $v$. There are 16 $VV$, 92 $Vv$, and 12 $vv$ individuals.
If I did this right, that means

$V = (16\times 2 + 92)/240 = 0.516$
$v = (92 + 12\times2)/240 = 0.483$

and then the expected frequencies are:

$VV = 0.516\times0.516 = 0.266$
$Vv = 2\times0.516\times0.483 = 0.498$
$vv = 0.483\times0.483 = 0.233$

which are different from the observed frequencies:

$VV = 16/(120) = 0.133$
$Vv = 92/(120) = 0.766$
$vv = 12/(120) = 0.1$

The book says that the population is evolving because the observed frequencies are different from the expected frequencies. What exactly does that mean though? How can you tell if the population is evolving if you don't even know the allele frequencies or the genotype frequencies of the next generation? The reason I ask is because I thought to disrupt the Hardy-Weinberg equilibrium means that the allele frequencies would change over generations, so could you say the population is evolving is you're only given the data for one single generation? I mean, you can get the same allele frequencies from both the expected and the observed genotype frequencies, so I don't know if the population can be said to be evolving. Maybe I'm understanding this wrong. Please help. Thank you all.
My related question is: Can you have no change in allele frequencies across generations, while in each generation, the observed genotype frequency is different from the expected genotype frequency. If so, does this still mean the population is evolving?

From Wikipedia, bold added by me:

In population genetics, the Hardy–Weinberg principle, also known as the Hardy–Weinberg equilibrium, model, theorem, or law, states that allele and genotype frequencies in a population will remain constant from generation to generation in the absence of other evolutionary influences. These influences include genetic drift, mate choice, assortative mating, natural selection, sexual selection, mutation, gene flow, meiotic drive, genetic hitchhiking, population bottleneck, founder effect, inbreeding and outbreeding depression.

Therefore, if you observe allele frequencies that do not match what Hardy-Weinberg would suggest, that means one of those other things is happening. Think of Hardy-Weinberg a bit like the "null hypothesis". You can't tell directly which of these is occurring, and multiple evolutionary influences can occur simultaneously.
We can think about a particular example, though. Let's say you have a case with a recessive trait where the phenotype of vv individuals makes them more likely to die. You would expect when you look at genotype frequencies to find them out of equilibrium with respect to Hardy-Weinberg: you'll find fewer vv individuals than you'd otherwise expect based on the overall rate of v alleles in the population.
It's important to consider sampling effects in your measurement, too. Your estimates will rarely match the Hardy-Weinberg frequencies perfectly just from noise in your sampling. You'll want some estimate of how likely it is that, assuming the population actually does follow Hardy-Weinberg, you'd get a deviation as large as the one you observe. That is achieved by performing a chi-squared test.

How can you tell if the population is evolving if you don't even know the allele frequencies or the genotype frequencies of the next generation?

You can tell because if the population isn't evolving (this statement means exactly that the allele frequencies stay the same) and mating is random then the allele frequencies would be equal to the Hardy-Weinberg frequencies. A bit like if you park your bicycle, go inside, and come back outside and see no bicycle there, you can infer that your bicycle has moved even though you didn't actually observe it moving. You may not be able to conclude whether it was ridden away, was blown by the wind, or picked up and placed in a truck and driven off, but just the fact that it's not where you left it provides information that something is happening besides the "null hypothesis" that bicycles left in a place stay there.

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What happens to eyelids when we feel sleepy?
Why is that whenever we feel sleepy, our eyelids start to feel heavier.

It is not like we can any time close our eyes and will immediately fall asleep. But, when we feel sleepy our eyelids automatically become heavy.

What causes heaviness of eyelids? Is it because of sleep hormone or something?

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Confused about the calculation of BLOSUM matrix
I am trying to understand how the BLOSUM matrix is calculated.
I found a very nice tutorial here: http://www.cs.columbia.edu/4761/assignments/assignment1/reference1.pdf
Also i read through the wikipedia article, and a book "Understanding bioinformatics" from Marketa Zvelebim and Jeremy O. Baum
After i thought i understand the method i tried myself with this "alignment" (i will refer to the variable as they are at the link i provided)
Q
Q
S

So it has only one column, and 2 type of amino acids.
The Count pair frequencies (C) for Q -> Q substitution is 1 (2*1/2) and for Q -> S it should be 2 (2*1).
To normalise it T = 3 as 3*2/2
The q values then seem to be q(Q->Q) = 1/3, q(Q->S) = 2/3
The p(Q) is then 1/3 + 2/3/2 = 4/6
p(S) is 2/3+1/3/2 = 2/6
The e(Q->Q) = (4/6)^2 = 4/9
e(Q->S) = 4/6*2/6*2 = 4/9
Thus
S(Q->Q) = round(log2((1/3)/(4/9)) = -1
S(Q->S) = round(log2((2/3)/(4/9)) = 1
So from this alignment it is more likely to substitute a Q to an S rather then Q to Q? This makes no sense for me.
What am i missing?

Ok I'll try to go through it myself and see what happens.
First, we get the pair counts: $$ c_{ij}=\begin{cases} \displaystyle\binom{n_i}{2} & i=j\\ n_i n_j & i\ne j \end{cases} \;\;\;\;\;\implies\;\;\;\;\; c_{11}=1, c_{12}=2, c_{22}=0 $$ Clearly, the normalizer is just $T=\sum_{i\geq j} c_{ij}=3$ and $c_{ij}\equiv c_{ij}^{(1)}$ since there's only one column. Then our pair distribution is: $$ q_{ij} = \frac{c_{ij}}{T} \;\;\;\;\;\implies\;\;\;\;\; q_{11}=\frac{1}{3},\;q_{12}=\frac{2}{3},\; q_{22}= 0  $$
Hence, the occurrence probability per residue: $$ p_i = q_{ii} + \sum_{i\ne j} \frac{q_{ij}}{2} \;\;\;\;\;\implies\;\;\;\;\; p_1 = \frac{2}{3},\;p_2=\frac{1}{3} $$
and the expected pair frequency:
$$ e_{ij} = \begin{cases} p_i^2 & i=j\\ 2p_ip_j & i\ne j \end{cases}\;\;\;\;\;\implies\;\;\;\;\; e_{11}=e_{12}=\frac{4}{9},\; e_{22}=\frac{1}{9} $$
Now we can finally compute the odds ratios:
$$ r_{ij} = \frac{q_{ij}}{e_{ij}} \;\;\;\;\;\implies\;\;\;\;\; r_{11}=\frac{3}{4},\; r_{12}=\frac{3}{2}, r_{22}=0 $$
Computing the log-odds $s_{ij} = \log_2( r_{ij} )$ gives the same values as you get.
So why is $r_{11} > r_{12}$? Well, notice they have the same normalizer, i.e. $e_{11}=e_{12}$. So the difference comes from the fact that $ q_{11} > q_{12} $, the empirical pair distribution. Essentially, because of the way we computed $q_{ij}$, we consider there to be only one pair QQ but two pairs of QS.
I would also note that sometimes results are a little unintuitive for very small samples (data sparsity throws off the statistics essentially).

Well I don't know whether this is right or wrong (I suspect right) but considering the irrelevant nonsense that gets up voted for other questions this surely deserves a +1.

This is what i thought, thabks for the confirmation!

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text:
In what cases can $k_{\rm cat}/K_{\rm M}$ be used to compare catalytic efficiency？
The ratio $k_{\rm cat}/K_{\rm M}$ is often used to compare the catalytic efficiency of enzymes, but this  paper has suggested that $k_{\rm cat}/K_{\rm M}$ should be abandoned for enzyme comparison. This confuses me. Should this ratio not be used in any cases, or can it still be used under some conditions, such as under low substrate concentration, $[\ce{S}]\ll K_{\rm M}$? If $k_{\rm cat}/K_{\rm M}$ is abandoned, how can we comprehensively analyze and compare the efficiency of enzymes? $k_{\rm cat}$ alone seems to only be applicable when $[\ce{S}]\gg K_{\rm M}$.

This appears to be an improved formulation of a previous question, in which case you should delete the original.


text:
What would be the growth rate in the following scenario?
In my textbook the exponential growth rate is described as
$$W'= W e^{rt}.$$
Here $W'$ is the final size, $W$ initial size, $r$ growth rate and $t$ time.
I wanted to have a clear mathematical understanding. So far according to what I understand, if the size is growing by 10% per unit of time, then the growth rate in the above function should be
$$r=\ln(1.1).$$
Am I right? Considering infinite resources.

So why don't we just write it as W λ^t? I have some faint idea that it's to represent that it's a derivative of itself and that it's change in value depends upon itself but I am not sure how that works exactly.

I've been doing some brainstorming to figure out why we write this function in "e" form, so please tell me if I'm right. It's just to make the calculations of rate a bit more standard (as the rate is changing because it depends on it's initial value, even though the percentage change is same). Without e we would have to find the proportionality factor for every individual percent change to figure out what the rate was. Right??

My comments disappeared, so I'll explain here:
There are two forms of the exponential growth model, in discrete time or continuous time.
A discrete-time model is most appropriate for a population that grows at discrete times in regular intervals, such as an animal that reproduces during a specific breeding season. We only consider t = 0, 1, 2, etc., and not the times in-between. The discrete-time exponential growth model is derived from N(t + 1) = λ N(t), where N(t + 1) is the population 1 time step after N(t) - that is, the population at any one time step is λ times the population at the previous time step. From this, we get N(t) = N(0) λ^t.
A continuous-time model is appropriate when growth happens continuously, without seasonality, etc. - perhaps a human population - or when time steps are negligible relative to the time period of interest. The continuous-time exponential growth model is dN(t)/dt = r N(t) - that is, the population grows at a rate of r times the population. By solving the differential equation, we get N(t) = N(0) e^(rt). Euler's constant comes up because of its unique property, d[e^x]/dx = e^x, so dN(t)/dt = d[N(0) e^(rt)]/dx = r N(0) e^(rt) = r N(t)
These two models are equivalent when r = ln(λ). (Note that the symbols r and  λ are commonly, but far from always used for these values.) So whether we choose to express the model in as N(t) = N(0) λ^t or N(t) = N(0) e^(rt) comes down to whether it is more intuitive to think about the growth rate in discrete time (λ) or continuous time (r).

Generally you can rearrange the exponential growth equation to get
r = ln(W'/W) / t.
So indeed if time t = 1 and W'/W = 1.1, then your growth rate would be
r = ln(1.1).
For example in microbiology frequently also the doubling time tD is used, thereby W'/W = 2 and growth rate r = ln(2) / tD.


text:
Help in identifying marine specimen?
I came across this specimen which I assume belongs toeither the phylum porifera or to the coelenterata.
Can someone please help me identify it?
The specimen was found in Ayodhya, India.

Please update your question by including where you found this specimen. Thanks

I am thinking it is the skeleton of a colonial large polyp stony coral. I am not certain, but I suspect a species of the Favia genus. Most likely one of these two species (see links for info and pictures):

Favia lizardensis
Favia rosaria

Looks quite similar. However, many stony corals appear rather similar when dead. Could you add additional information in support of your answer?


text:
T-cell "exhaustion" in patients with Covid-19
Following up on my previously asked question, which asks how COVID infects T-cells in immune system.
Here's a study that shows that recovered covid patients have "exausted t-cells". To quote:

T cell counts are reduced significantly in COVID-19 patients, and the
  surviving T cells appear functionally exhausted. Non-ICU patients,
  with total T cells, CD8+T cells CD4+T cells counts lower than 800/μL,
  300/μL, and 400/μL, respectively, may still require aggressive
  intervention even in the immediate absence of more severe symptoms due
  to a high risk for further deterioration in condition.

Is this normal for other coronaviruses (like the common cold)? Also is it possible that this immune damage can be a long-term effect, similar to HIV?

It's not a flu-like virus! Symptoms may be similar, since they are not specific, though flu doesn't cause interstitial inflammations and pulmonary embolism!

I edited this so that it's more clear. In a previous question people said that it's normal for coronaviruses to infect T-cells. By "flu-like" I was referring to other coronaviruses like the common cold that spread through airborn droplets like the flu.


text:
Automating figure-making with PyMOL
What I want to do
I have a list of almost 200 PDB entries that I need to visually inspect. Luckily, the thumbnail image displayed on the PDB website (examples with RCSB or PDBe for a well-known structure) is enough to figure out whether I want to exclude an entry for subsequent use of the resulting cleaned-up list of entries.
Now, looking up these entries one by one on the website is easy to do, but is very slow and error-prone. I need to be careful copy/pasting accession codes, and it takes a lot of time to go to my list, copy an accession code, go to the website, paste, search, and have my answer once the result page has loaded fully.
In other words, I'm lazy and I think my time would be better spent doing something more interesting than copy/pasting 200 times.
Attempt at a solution
I figured it shouldn't be too difficult to use PyMOL with a bit of scripting to automatically fetch each entry and generate a small image (faster than ray-tracing, which I don't need for this quick inspection). Here is my attempt:
pdb_entries = ['1m18', '1eqz'] # The real list has almost 200 entries, but 2 are enough to test the script

for entry in pdb_entries:
    cmd.fetch(entry, async = 0, path = "./pdb-files")
    filename = "./pdb-imgs/" + entry
    cmd.png(filename, width = 600, dpi = 100, ray = 0, quiet = 0)
    cmd.delete(entry)

Problem
When I save this script as fetch-all.py and run it from within PyMOL with the command run fetch-all.py, the structure files are correctly downloaded in the ./pdb-files directory and images with names matching the PDB accession codes are generated in the ./pdb-imgs directory. So, it seems everything works. Except these images are just black, not showing anything of the structure.
I tried to add cmd.center(entry) and cmd.show("cartoon", entry) before saving the PNG files, but it did not change anything. I also looked up the help page for cmd.png(), thinking I was missing something, but I couldn't find anything I would be obviously missing there.
Any idea? Thank you all in advance.
Edit: I forgot to mention that this problem of black images happens with the latest PyMOL (2.2.0) and also with an older version (1.7.4).
Edit: when I run the commands inside the loop one by one at the PyMOL prompt, everything works fine, I get images correctly saved. It seems it doesn't work in the context of the loop only. I double checked the loop syntax and it looks fine to me (but I might be missing something; I don't know python very well).
Edit: I also tried cmd.save() instead of cmd.png(), but it did not solve the problem.

Could it be that as all your structures have different x, y and z coordinates. Therefore whereever pymol is centered  it isnt in a position where the protein is?

Therefore could be fixed by aligning all the structures so they all appear in the same x, y and z range

I tried to add cmd.center(entry) in the loop that processes each entry, to make sure that each entry gets properly centered before saving a picture. That didn't solve the problem.

Have you tried using cmd.orient ?

I just gave it a try, it did not solve the problem of the black images.

Were you able to fix this problem

I ended up using ChimeraX instead, which I found easier to use overall. I abandoned the idea of generating pictures and browsed the models interactively in ChimeraX (this is slower than looking at pictures, but being able to rotate the model was useful in some cases).

I’m voting to close this question because it is not concerned with a biological problem.


text:
do tumour cells begin with abnormal characteristics?
At what point in the cell cycle do cells start to become tumorous? Do they have abnormal characteristics to begin with; if so what are they?

No, I recommend you look at the stages of tumour development. There are both environmental and genetic factors which contribute to this.

Cancer cells don't start to become cancerous at a specific stage of the cell cycle; you will find that while uncontrolled proliferation is a hallmark of cancer, different cancers acquire alterations in different phases of the cell cycle. BRCA-deficient cancers for example have a compromised G2-M checkpoint [1], while Rb deficient cancers have a compromised G1-S [2] checkpoint in the cell cycle.
The cell cycle is simply a property of proliferating cells and the same broad phases of the cell cycle are universal to both normal and malignant cells.
As for when abnormally growing cells actually become a cancer - this has nothing to do with phases of the cell cycle, and everything to do with the ability to break through the basement membrane of the original site (indicating the potential to be invasive/spread), because the ability to invade is a hallmark of cancer [3]
References
[1] http://cancerres.aacrjournals.org/content/67/13/6286
[2]  https://www.ncbi.nlm.nih.gov/pubmed/16936740
[3]  https://www.ncbi.nlm.nih.gov/pubmed/21376230

In some ways this is simply a function of how we define 'tumorous'. We can trace the phylogeny of tumors in the same way we trace that of species, using DNA sequencing. In some tumors one sees a clear causative mutation at the root of the tree, so for these we might say that there was a single mutation which caused cells to become tumorous. Mostly though things are not so clear.
For instance, it's now believed that in many cancers, the rapidly dividing cells are descended from more slowly dividing 'cancer stem cells'. These cell cancer stem cells tend to survive therapies that target more rapidly dividing cells, such as radiotherapy or many chemotherapies. In such cancers we could talk about a mutation by mutation transition from non tumorous to tumorous cells.


text:
Mice that have been reconstituted with geneX-transduced cells
I read this article in the following:

... Mice that have been reconstituted with geneX-transduced cells

Now, someone who is not familiar with animal sciences, what does this exactly mean? It is in the context of hematopoietic stem cells. I have two possible interpretations as a result:

An Hematapoetic stem cell with geneX was successfully transplanted into the mouse
The mouse cells (from the embryo) were engineered to contain geneX, and hence the mouse has been reconstituted with geneX transduced cells

Are any of these correct? If I am wrong please let me know!

Your second interpretation is correct.  The mice have been reconstituted from stem cells that were "transduced" (using a viral vector) with a construct containing the appropriate geneX. The link gives comparison of transduction vs. transfection.
https://www.vectalys.com/blog/what-are-the-differences-between-transfection-and-transduction-222


text:
Is Plasmid pGLO a stable or unstable plasmid varient ?
Is plasmid pGLO a stable or unstable plasmid varient?

What research have you done before asking it here?

What do you mean by an unstable plasmid?

The GFP protein is an unstable variant

The pGLO plasmid uses a stable version of GFP. This is stated in the pGLO education series protocol, and wiki.
I could however, engineer to be unstable if wanted.
https://en.wikipedia.org/wiki/PGLO


text:
Picking up Plasmid DNA using Nanodrop, but not using Electrophoresis
I am currently trying to see if 2 bacteria contains plasmids or not. I had used promega plasmid extraction kit on the bacteria. I ran a gel electrophoresis (.8% gel) with the extract product, however its not showing the presents of any DNA. I also ran the product through Nanodrop (2000) and these where the concentrations I got:
1 : 6.5 ng/ul & 260/280 = 1.56
 2 : 49.4 ng/ul & 260/280: 1.96
Based on experience, I want to say #1 contains no plasmids, however #2 may have a small concentration of plasmids. Please give me your opinion on what to conclude. Also, if there is another test I can try to do to confirm plasmid presents or not, please let me know.

If your samples are contaminated with proteins (which can happen), then the resulting OD 260/280 gets higher and can falsely show some nucleic acids. If you run a few microliters on a gel and don't see anything, I would be suspicious. The concentration is sufficient to see something.

The amount of DNA is what matters when running the gel, not just the concentration. What volume of each sample did you run on the gel? Use the concentrations determined above to see how much DNA should have been there. If the hypothetical amount looks good, and you don't see anything on gel, I'd say there isn't any real DNA. As far as testing for presence of plasmid, you could try re-transfecting your suspicious sample into other bacteria, since you don't need many copies of the plasmid to get some good colonies.

This could be. I believe that the nano drop shows protein contamination. I am going to try to concentrate the plasmid DNA, and increase volume for gel. If I can not find plasmids, then I am going to assume that there are no proteins.

I don't know how much culture volume you used, but the amounts you have look more like a medium/high copy plasmid. I don't know if that's what you'd expect for a natural plasmid.
In addition to the other answer, it could also be sheared genomic DNA, cosmid (that would run too high on your gel to see), or maybe just nucleotides (if you messed up the prep in some way).
Another strategy to figure out if your strain has plasmids depends on what else you know about these strains. If you don't know that much: do 16S sequencing, it might be a known strain with known plasmids. If you already know it's a newly discovered strain: sequence the whole thing. Even with relatively low coverage the presence of plasmids will be quite clear. It's a bit more expensive than a plasmid prep + gel, but you'll also get a lot of other information about the organism.

Having a positive reading on the Nanodrop but no band on the gel can be due to a number of different issues (not exhaustive):

An incorrectly zeroed sample
Incomplete staining of the gel
Contamination of the sample
Human errors in loading of samples

You should try to exclude these possibilities by positive controls in order to find out the true reason behind the inconsistent results.

Depending on how you followed the protocol you could theoretical also have bacterial genomic DNA in your sample. If you follow the protocol strictly this should not happen, but if so, you would be able to measure the genomic DNA. In the agarose gel, however, it would usually be stuck around the top since it is too large to migrate.
Vortexing the sample before loading the column could cause this, for example (see Promega PureYield Manual).

Geo - if you could, please add some references/citations to your answer - I know at times it may be hard to reference something that may be common knowledge to you, but we appreciate it very much on this site. Thanks for your contribution!

I would recommend switching up your PCR cloning program. Often, even after purifying DNA and RNA in conjunction with great care PCR can be the bottle neck for Gel Electrophoresis, as well as ensuring the quality (depth, void of blunt edges, etc) of the agarose gel- 49 ng/ul is not enough for  progression. If all that still doesn't improve your case then it may be an issue with cultivating bacterica cultures -after purifying plasmids from bacterial clones vials should appear very cloudy (8-16 hrs was my go to time period), futhermore make sure to avoid light exposure on to the agarose gel as well as keeping it clean and submerged under cold PBS before inserting your buffered DNA (I used 10 microliters - but it tends to vary depending on the thickness of your gel) Also, when I would use a higher voltage during gel EP I would have a higher chance of having vertical streaking though it may be faster.
If nothing works... then I can only allude to possible human errors. Last thing! I hope you clean the nanodrop before you take count, having protein in your product can be a result of that. good luck! bacterica need love too!

Christopher - if you could, please add some references or external citations. We always appreciate it on this site. Thanks for your contributions!


text:
What is a "NuMA protein"?
I just got out of a lecture on mitosis and this part was incredibly confusing. After searching my textbook it doesn't cover it that well either.
The professor mentioned a NUMA protein that is released after NEBD, and that it helps form more Microtubules. I don't really understand how it does this.
Is it just a protein that sits in the pericentriolar matrix and encourages microtubule nucleation? Or is it a a kinase that deactivates some GAPs that destabilize microtubules?
Also, if you would like, there's also this study question I feel is sort of related: What are two proteins located in the pericentriolar matrix that encourage microtubule nucleation?
I feel like one of them is the y-tubulin ring complex. Would the other be NUMA?
Thanks for reading.

you should ask your professor if you have doubts about a lecture. you can read about NuMA here


text:
Is it safe to look indirectly at the Sun?
I.e., is exposure of sunlight onto the peripheral vision less damaging than exposure onto the fovea?

Behar-Cohen et al. notes the periphery of the cornea has a focusing effect:

UVR incident from the periphery is refracted into the eye, and due to the focusing effect of the cornea, UV radiation is on average 22-fold stronger at the nasal limbus, which is the typical site for pterygium and pinguecula. Moreover, UV radiation is on average eight times stronger at the nasal lens cortex, the typical site for cortical cataract.

Authors note this in further papers as peripheral light focusing. Essentially, the light is refracted from the periphery of the cornea into the anterior segment of the eye. By this notion, I'd say indirectly looking at the sun is no less damaging than directly looking at it (given there are no other restrictions, such as special lenses, etc.).

Interesting that they only say, 'never look directly at the Sun,' then


text:
Are flowers / flowering plants vital to all life on Earth?
Not a biology student so forgive me if this is a very basic question. Are flowering plants (angiosperms) vital to all (or most) life on Earth?
In other words, if flowering plants disappeared, would all (or most) life on this planet be gone as a result?
To give a bit more context, I'm arguing with a friend who claims (from a biblical perspective) that without flowering plants life couldn't exist.

I'm curious, what is this biblical argument that life can't exist without flowers? (if you know)

Angiosperms -- that is, flowering plants -- only evolved relatively recently on an evolutionary timescale, about 125 millions years ago. So for most of the history of life on earth there have been no flowering plants. Thus it seems highly likely that if angiosperms were to suddenly disappear, life on earth would continue. There might be a massive disruption of current terrestrial communities, but it's hard to imagine this resulting in total global extinction. In particular it seems unlikely that ocean communities would be undermined by the loss of flowering plants.

To put that in context, the earliest life dates from at least 3 billion years ago (from stromatolite evidence). Of course, some Christians may dispute that the earth was around even 125 million years ago, so this argument probably doesn't hold in the face of such claims.


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What are the applications of predicting the structure of proteins?
Protein molecules are very important as they are used for catalyzing almost all the chemical reactions in the cell, regulation of gene activity and provide cellular structure.
However, in predicting the structure of protein I haven't been able to find many use cases. In the omicsonline I was able to find a paper on Computational Methods for Protein Structure Prediction and Its Application in Drug Design:

Most modern drug discovery projects start with protein target identification and verification to obtain a verified drug target. For structure-based drug design the three-dimensional structure of the protein needs to be determined experimentally by using either x-ray crystallography or nuclear magnetic resonance (NMR) spectroscopy.

Therefore, since proteins are vital for maintaining the body are there any other use causes for predicting the structure?

To answer your question specifically in the context of drug design: it would of course be great if we could rely on protein structure prediction for drug design, but structure prediction is still not completely accurate. Drug design projects are expensive, so it is often wiser to spend the money on experimental structure determination first, because it will provide a more reliable basis for structure-guided drug design (the alternative would be to spend the money on more failed attempts at designing a good drug, because these attempts are guided by an inaccurate predicted structure).
Also, experimental structure determination is valuable for prediction, because it increases the library of known structures for homology modeling.

Just off the top of my head, protein structure is very important for determining how proteins interact, for example determining molecular pathways or viral capsid structures. Knowing the structure of a protein can help scientists better understand what it does and how it does it. By looking at protein structures, scientists can determine exactly how a different allele for a gene causes changes in an organism's phenotype at a biochemical level. Predicting sites within a protein's structure that have an enzymatic function allows scientists to predict how various factors effect it's function. Differences in temperature or pH can have an effect on protein structure as well, and understanding how a proteins changes based on pH or temperature changes can help scientists determine how a protein functions in its specific environment.

protein structure prediction is still in its infancy hence the lack of examples, the benefits are immense however, in proteins structure = function, and as you said proteins catalyze or make up nearly everything in body. We can find out the structure unreliably but predicting the structure would allow us to build proteins to order becasue predicting the structure from sequence means predicting protein folding, which is in many way the holy grail of modern biology. Being able to do it reliably and with more complex proteins would allow us to construct proteins to performs specific tasks instead of hoping to stumble across one that does it then just copying it.


text:
Laemmli-SDS-PAGE problems
I did Laemmli-SDS-PAGE for my Ammonium sulphate precipitate but I had very weak band and have very weird part at the end of gel. Please help me to solve that problem. Thanks

Did you remove the ammonium sulfate by dialysis or similar?

No, I did not. I precipitated by AS and used PBS to dissolve, added buffer and loaded on gel

Your gel looks just fine to me. It simply indicates that your ammonium sulfate precipitate contains massive amounts of this small protein at the bottom (whatever this is). The upper bands look fine, the amount loaded is appropriate: they are not overloaded like the bottom one, but not too faint either. The fact that your first and last lanes look a bit smeary is because of the massive amount of the small protein (lanes do look smeary when overloaded this much). The bottom band is definitely really some protein, and not a staining artifact; I have seen this shape before with purified proteins overloaded on SDS-PAGE.
One thing you could do, assuming you still have enough of these samples, is to run a complementary gel with 100 times less material. Dilute by a factor of 1/100 in 1X Laemmli denaturation buffer, and load a new gel with the same volume as for this one. You won't see the upper bands anymore, but you will have a much cleaner bottom band that you will hopefully be able to assign an apparent molecular weight to (or even cut out for mass spec identification; wear gloves and use clean tools if you do that, otherwise they will mostly detect keratin from your skin).
Assuming the point of this gel was to check the outcome of ammonium sulfate precipitation used as a purification step, well obviously it didn't work well to separate all these bands, and you might need to repeat and try different concentrations. But if the bottom band is the one you are trying to purify, then you were successful (the top bands represent less than 10% contamination, I would say by eyeball estimation). If you're interested in one of the top bands, then it clearly didn't work.
Is one of these lanes the supernatant? If so, you also have massive amounts of the small protein there (it's in every lane). Such a high enrichment looks suspicious: could that be purified lysozyme that you added during the purification procedure to help break down bacterial cell wall? (assuming you're trying to purify a protein over-expressed in E. coli).

Thanks you for your respond. I would like to determine the molecular weight of the bottom band, because when I overlaid the half of gel with pathogens, the place of inhibition zone is the bottom of the gel (the second lane, beside ladder) . I will try to dilute the buffer as you suggest. Thanks

OK, so if you're interested in the bottom band you can definitely dilute (I suggested by 1/100, but you might need to try a few different dilution factors until you get a reasonable band). You can also use a denser gel, to make this band migrate slower (in this gel, it looks like this band is in the migration front). What was the percentage of this gel? Also, if the answer helped you, please mark it as accepted. :-)

Hi, I tried to dilute it to several different dilutions but it was not different as before, I did not get a reasonable band. The percentage of this gel is 4- 20%. I think maybe the size of this compound is smaller 11KDa (because I used Color Protein Standard marker, 11-245 KDa) or this compound is not protein, like lipid


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How to choose right volume of DNA and SYBR Green
I'm testing fluorescence level of a sample having dsDNA, SYBR Green and milli q water but I'm facing difficulty in choosing right volume of dsDNA and cons level of SYBR Green.
Details of sample: dsDNA:1000 nano mole, SYBR Green:1X (1000 times diluted from original stock)
Sample_1: dsDNA (3ul) + SYBR Green (5ul) + milli q water (27ul)
Sample_2: dsDNA (3ul) + SYBR Green (5ul)
Result: In first sample I haven't got any signal but in second case I got( But I want 5 times higher then what I got).
Please tell me how it work and how these things depends on each other. I'm not a biologist if possible suggest me resource as well.

I had to look it up online, but I eventually found here an important information: the detection limits of SYBR Green.

dsDNA at 300nm transillumination: 60 pg
  Oligonucleotides at 300nm transillumination 1-2 ng

You should first check the size of your DNA of interest, and from its size figure out its approximate molecular weight (for example using this online calculator). Then, you should check that 1000 nmoles of your DNA corresponds to a mass greater than 60 pg. If not, you should increase the amount of DNA in your assay accordingly until the mass introduced in the assay exceeds the detection limit of SYBR Green.
Now, do you know the concentration of your DNA? In other words, how did you choose to use 3 uL of DNA? How do you know that these 3 uL contain enough material? (1000 nmoles as in your protocol, or better, that this is higher than the detection limit in terms of mass).
Also, did you use the 1000X stock solution of SYBR Green without intermediate dilution? If so, the final concentration of SYBR Green is much larger than the final 1X you want, in both of your samples:

sample 1: 5 uL of 1000X SYBR Green in a total volume of 3 + 5 + 27 = 35 uL means that the final concentration is 1000X * 5 / 35 = ~143X.
sample 2: 5 uL of 1000X SYBR Green in a total volume of 3 + 5 = 8 uL means that the final concentration is 1000X * 5 / 8 = 625X.

What you need is either use 1 uL of 1000X stock solution of SYBR Green + whatever amount of DNA that will be above the detection limit + buffer to adjust the volume to 1000 uL final (which will bring the SYBR Green to 1X final), or make an appropriate intermediate dilution of SYBR Green so that its final concentration is 1X by following the sample preparation as described in your question.
The sample preparation you describe here yields a composition that is way off the one suggested by your protocol, at least for the SYBR Green. Until you try again with proper final concentrations, do not conclude that it didn't work. :-)
Good luck with your project.


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Cutting of DNA strand
Would EcoRI cut the following sequence??
5'CTCGAGTTCGAG3'
3'GAGCTCAAGCTC5'
What is the logic of cutting? Please explain how restriction enzymes work.

No, it will not cut it, because EcoRI recognizes the following palindromic sequence: GAATTC, and only if the whole sequence is present it will cut it at particular site. You can find these information for all restriction enzymes at sites of the providers, for example at New England Biolabs site https://www.neb.com , just enter the name of the enzyme you are interested in.


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Blackgram leaves. SDS page. Phosphate buffer method

I am doing protein analysis in blackgram leaves by using phosphate buffer method. But I cannot get proper bands on my SDS-PAGE. What should I do to get proper bands?

Can you please exactly describe what you have done? At the moment this is not possible.

You should de-stain your gel longer. I can see at least one thick band at the top, but without de-staining longer you are probably missing other fainter bands.
Also, please explain what you mean by "proper bands". The top one looks fine to me. There might be others but we cannot see them.

There are other bands below; definitely destain. But perhaps the OP is referring to the frowning?

In protein analysis lots of bands can be seen in the gel bt I got only one band. That's why I am asking what should I do? Can you send a proper protocol to me. My mail I'd mohanlal0606@gmail.com. please help me by sending detailed protocol.. Thank you all for your comments..

@Mohanlal seeing a single band might be a good result, depending on what you are trying to achieve. For example, if your goal is to purify a protein, it's very good news when a single band shows up on an SDS-PAGE (assuming this is the correct protein, of course). I don't think you need to change anything to your electrophoresis protocol, except de-staining longer. You might need to change whatever you are doing before the SDS-PAGE if this single-band result is not what you want.


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The total numbers of mutations on different branches of gene tree are independent or not?
I was reading Tajima's 1989 paper on his test for neutrality.
Tajima, Fumio. "Statistical method for testing the neutral mutation hypothesis by DNA polymorphism." Genetics 123.3 (1989): 585-595.
Here is the question:
Suppose we have three sequences labelled $C,D,E$, and their genealogy follows $\{\{CD\}E\}$  (i.e. $C$ and $D$ coalesce first before they coalesce with $E$). Let $B$ be the most recent common ancestor of $C$ and $D$.
Now, let the random variable $k_{ij}$ be the number of nucleotide differences between sequence $i$ and sequence $j$, then Tajima shows that $k_{BC}$ and $k_{BD}$ have a non zero covariance.
But aren't mutations in the branch $BC$ independent from mutations in the branch $BD$? I was thinking that the total numbers of mutations in the brand $BC$ and $BD$ are two independent identically distributed variables, so $k_{BC}$ and $k_{BD}$ are independent, then why they have a non-zero covariance?
============Update=============
Now I have some basic ideas, but haven't worked out the full answer.
Tajima's definition of $k_{ij}$ is neither independent of the sample size, nor for a fixed coalescent time. (See his 1983 paper: Tajima, Fumio. "Evolutionary relationship of DNA sequences in finite populations." Genetics 105.2 (1983): 437-460.)
For example, in a sample size of 3, if you pick 2 individuals, and condition on that the two coalesce first, their coalescent time will follow:
\begin{align*}
\mathbb{P}(t=T)=p(T)=\frac{3}{2N}e^{-\frac{3}{2N}T}
\end{align*}
Now conditioning on a fixed coalescent time $t$, the number of mutations under the infinite sites model in each branch either from $B$ to $C$ or from $B$ to $D$ will follow a poisson distribution with parameter $\mu t$, where $\mu$ is the mutation rate per sequence per generation. Let this poisson random variable be $\xi_t$ in branch $BC$ and $\eta_t$ in branch $BD$. Then
\begin{align*}
k_{BC}=\sum_{t=0}^{\infty}\xi_tp(t)\\
k_{BD}=\sum_{t=0}^{\infty}\eta_tp(t)
\end{align*}
If we only consider the partial sum of the above series,
\begin{align*}
k_{BC}^{(n)}=\sum_{t=0}^{n}\xi_tp(t)\\
k_{BD}^{(m)}=\sum_{t=0}^{n}\eta_tp(t)
\end{align*}
then $k_{BC}^{(n)}$ and $k_{BD}^{(n)}$ clearly have a zero covariance, because $\xi_t$ and $\eta_t$ are independent poisson variables, hence
\begin{align*}
\mathbb{E}(k_{BC}^{(n)}k_{BD}^{(n)})&=\mathbb{E}(\sum_{t=0}^{n}\xi_tp(t)\sum_{t=0}^{n}\eta_tp(t))=\sum_{i=0}^{n}\sum_{j=0}^np(i)p(j)\mathbb{E}(\xi_i\eta_j)=\sum_{i=0}^{n}\sum_{j=0}^np(i)p(j)\mathbb{E}\xi_i\mathbb{E}\eta_j\\
&=\mathbb{E}k_{BC}^{(n)}\mathbb{E}k_{BD}^{(n)},
\end{align*}
so their covariance is zero.
But as $n\to+\infty$, how $k_{BC}^{(n)}k_{BD}^{(n)}$ converges to $k_{BC}k_{BD}$ is questionable. It will not uniformly converge to $k_{BC}k_{BD}$ because otherwise we can first computing the expectation then take the limit, which gives us a zero covariance. Tajima didn't explicitly show us how he computed the covariance by summing together three infinite series (Line 7, Pg. 448, 1983's paper). I tried directly working on that series but failed in the last sum. His result is correct, though, I hope someone can give some hint on why there is inherent correlation between these seemingly independent random variables.
=======Update: a simple explanation has been posted==============

Could you please highlight where in the paper (which equation) is it claimed that $cov(k_{BC}, K_{BD}) ≠ 0$?

If the ancestor $B$ is not considered as a single sequence but as a population of sequences, then if B has many polymorphic sites $S$ and a high average heterozygosity $\pi$, then drift will have an important say in the values $k_{BC}$ and $k_{BD}$ and both values will be large. If the ancestor $B$, has $S=0$, then the only contributors to $k_{BC}$ and $k_{BD}$ are new mutations and hence both values will be small.

Hi Remi, the equation is at Pg. 586 at the middle right panel, roughly 5 lines above Equation (15). It is originally derived from His 1983 paper: Evolutionary relationships between DNA sequences in finite populations

But the coalescent process only considers the relationship among sampled individuals, so I think each label should be a unique sequence.

Hmmm.... Yes, I now agree with you. The graph makes it pretty clear that 'B','C', 'D' refers to individuals sequences. Would be worth going into this 1983 paper. +1 in the meantime. Good question!

Here is a simple answer to my question. The reason two numbers of total mutations accumulated in two divergent branches are not independent of each other is because they experience the same amount of coalescent time.
While the stationary Poisson mutation processes are independent from each other as long as they happen in different branches of a genealogy, they are likely to produce a similar number of mutations if two such processes happen together for a same amount of time. Thus, the non-zero part of the covariance between $k_{BC}$ and $k_{BD}$ is not from mutation processes themselves, but from the shared coalescent time $T$.


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Pits and Pores in Vascular Tissue
What purpose is served by the pits and pores in xylem and phloem cells? I cant find what purpose that these components serve in the vascular tissues in plants

Welcome to Biology S.E.! If you need additional assistance with the site, please visit The Help Center.

Welcome! Could you provide a reference to where you have encountered these features? A picture might help too. Good luck.


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Find the length of DNA sequence?
Given $N=5\times10^3$ and mutation rate is $\mu=10^{-5}$ per site, find the length of a DNA sequence so that the probability of mutation occuring M, is greater or equal than 0.95.
Is there a method or a formula for this type of calculation?

Welcome to Biology.SE. Homework (or homework-like) questions are off-topic on Biology unless you have shown your attempt at an answer. For more information see our homework policy.

Hint: Try to compute the probability that a mutation occurs at a single site (in a single generation). Then compute this same probability for 100 sites. And from there you should see what is going on.

is $n$ the population size? We typically use $N$ for the population size. Also, I would assume that $\mu$ is the per site (per base pair) mutation rate. Finally by probability of mutation, I would assume it is meant probability of 1 or more mutations, just like the formula is suggesting if $M$ means the number of mutations.

I just realized there are two ways to understand the question. By mutation do you mean mutant allele or do you mean mutational event (see here)?

I mean by an event , yes

So, you don't need any a priori about the sequence. The probability of a mutation to happen at a given site on a given haploid is $10^{-5}$. What is this probability for the whole population (don't forget individuals are probably assumed to be diploid)? You can then compute this probability for 100 sites and you'll understand the pattern. Does it make sense?

Ok, so for each site: $P(M>0) = 10^{-5} $ meaning that for $N=5000$,  we get $P(M>0) = 5000*10^{-5} = 0.05$ for one site?

I am really not sure how this method would allow us to find the length to ensure $P(M>0)\ge 0.95$. I know that $\theta = 2 N \mu $, which I think may be used so that $\theta=0.1$ per base. But i don't get how to answer this in terms of the actual question

Ok, good! 0.1 is correct. (0.05 would be correct if assuming haploidy). Now for 100 sites? And then just ask for how many sites is the probability 0.95

The mutation rate per haplotype per site is $\mu = 10^{-5}$. Assuming diploidy and a population size of $N=5000$, the population wide mutation rate per site is $10^{-5} * 5000 * 2 = 0.1$.
$0.1$ is hence the probability that a mutation occurs a at a given site (in the whole population). For 10 sites the probability that a mutation occurs at at least one site is $1 - (1-0.1)^{10} = 0.65$.
The probability we are aiming for is 0.95. So let's write the equation
$$1 - (1-0.1)^{x} = 0.95$$
, where $x$ is the number of sites we are looking for. You just have to solve for $x$ now and round up to the larger integer.

Ok, so this is in terms of the number of sites, so to calculate the length of the sequence in terms of base pairs, would we multiply by $10^3$ say? I am trying to read up on base pairs and this is what i gathered, but i may be mistaken. (Otherwise this answer is perfectly fine, i just see in many literature the lengths are usually given in bp?)

A site is a bp. The number of $x$ is the length of the sequence (in sites of in base pairs) that you were looking for. If you want to express the answer in kilo bp, then you can divide the result by $10^3$ but I would not recommend it. $x$, once rounded, is your answer

Ok! I am comparing this to mitochondrial dna which is generally 16,500 bp in length, so solving the above equation of 29 seems very unusual.. (ref: https://www.ncbi.nlm.nih.gov/m/pubmed/26798311/)

The answer depends upon the question. You typically don't attempt to answer everyday the question "what is the sequence length which probability to be hit by a mutation in a given generation is 0.95?". I don't think you should expect from yourself to have much intuition in the answer and therefore there is nothing unusual to be found. I don't understand why the length of the human mtDNA matters here.

Only because for instance the mutation rate (per site) of mtDNA is also generally $\mu  = 10^{-5}$ so i just wanted to compare

oh ok. Sure... so yes, a lot of mutations happen on mtDNA every generations (even more so if you consider 7 billions people).


text:
How to quantify the strength of social bonding between the individuals of a colony? How to compare the strength of social bonding across the species?
Consider two different species of honey bees. If I have to compare the strength of social bonding between the two species i.e. in which species, social bonding between the individuals is stronger than another.
Is there any methodology to do the comparison of the strength of social bonding across the species?

I don't know about bees, but in identified animals such as monkeys or river dolphins, scientists can count how many times two individuals are seen together in the same group. You still can mark individual bees, but that seems a lot tougher to do.

@Rodrigo I suspect it's actually easier to do per individual, you just need to do a lot more individuals... :P

@arboviral Then you would count how many times each individual is seen alone or with others? Seems easier, but it depends on the definition of "social bonds".

@Rodrigo yes, I'm not sure how that would work - I mean, all bees regularly visit the hive, so in that sense they're always "with others". But the marking per se wouldn't really be a problem - in fact, it's probably easier than for a lot of other insects just because queen marking is so useful there are commercial products and established techniques for marking individual bees, like numerical labels that glue onto the thorax (https://www.mannlakeltd.com/queen-marking-numbers).

@arboviral Yes, the problem is to monitor some bees in hives with up to hundreds of thousands of individual bees.

Thank you, Rodrigo, arboviral. Observing one colony and interaction between individuals is difficult, but could be possible. What about comparing two different species? @Rodrigo, could you please send the link of the papers? (I will also search the work done in other taxa)

I have been thinking on this. Can we consider the effect of isolation of various group sizes - (1Individua, 2, 4,  ......., 100, ... individual) from colony and see the relative change in the aspects like behaviour, life span, social network?


text:
Banana peel bruises after a hit
When a banana fall on the floor or get a certain impact, the impact location gets darker (it bruises) after a certain amount of time. The darkening affects the peel as well as the fruit itself. Why does this happen?

Related: https://biology.stackexchange.com/questions/16523/what-exactly-makes-bananas-go-brown

@LinuxBlanket Thx :) I saw that but, I was wondering what is happening when the banana fall and not only the normal ripening process. From what I have read online, it seems that when a certain pressure is exerted on the peel, the cells are damaged and they release an enzyme that will make the ripening faster. Nevertheless, I am waiting for a technically detailed answer describing the process.

Fruit is discolored in the first place because, when certain compounds are altered (oxidized), they turn brown.
In a banana (and most fruit) those compounds are phenolic compounds, and polyphenoloxidase is the thing that oxidizes them.
In an undamaged banana, the phenolic compounds and the polyphenoloxidase (that can turn them brown) are physically separated.
When the banana is crushed/dropped/etc, the structure of the banana is damaged so that the phenolic compounds and the polyphenoloxidase come into contact.  The polyphenoloxidase oxidizes the phenolic compounds, and you get that brownish color.
A nice summary (which links to a more detailed reference) can be found here: https://www.scientificamerican.com/article/why-does-bruised-fruit-tu/


text:
Is there a point in our life when ALL the atoms from our childhood's body get replaced?
The evolutionary biologist and Oxford professor Richard Dawkins makes on his TED talk the rather interesting suggestion that we human beings, are in nature more similar to a wave than an object. In the sense that we are a persistent structure travelling trough a constant flow of atoms that only transiently form part of our body.
Citing Steve Grand and his book Creation: Life and How to Make It, he argues that the body of a grown up man doesn't preserve any of the atoms that formed his body by the time of his earliest childhood's memories.
Is that fact widely accepted by the scientific community?
I know that this question is a duplicate of Are all body atoms really recycled several times during a life? (or this one), however I want to put the emphasis on the atom constituents of cells, as the accepted answer there doesn't fully answer that question.
Following those answers and the references within, a consensus seems to exist that most of the body is in constant renovation, with the exception of the cells in the cerebral cortex and the inner lens of the eye, whose cells persist trough life. But to what extent cellular metabolism replaces individual constituent molecules and atoms throughout those cell's life? Is that true also for DNA? What about mitochondrial DNA?
Are the processes that repair DNA damage operating only in damaged areas? Or do they constantly "repair" also healthy areas leading to the constant replacement of the atoms that constitute the DNA?
Others argue that tooth enamel does not regenerate or get replaced, been that what allows carbon dating to be accurate to the year of birth plus or minus a few months. However, as my question begins with a philosophical issue, I'm mostly interested in the atom constituents of the cerebral cortex. I am in particular aiming to figure out if we are more precisely described as a brain travelling through time or just a wave in a sea of atoms.

What defines childhood? Anytime before puberty?

@Sudachi Considering the context of the example I've pointed, it would be any age you can preserve memories of in adulthood. So I would guess from 3-4 years of age onward.

Is there a point in our life when ALL the atoms from our childhood's body gets replaced?

No. There is no point where ALL atoms have been replaced.
This has been shown by tracing distinct carbon isotopes in DNA [which reflect upon atmospheric 14C at the year of their generation (which reflect the extent of human atom bomb testing)]; Bergmann et al. 2009, Science: Evidence for Cardiomyocyte Renewal in Humans.

for your other questions:

Is that fact widely accepted by the scientific community?

The phrasing of Richard Dawkins, which you also list, uses "more similar" (rather than the absolute ALL). These words provide quite an accurate approximation to express that  most building blocks of cells undergo a constant renewal. The choice of words of Grand seems a bit unlucky, if read out of context.

Are the processes that repair DNA damage operating only in damaged areas?

As a first approximation: yes. For instance the image below shows the localized detection of H2A.X histones upon damage at the DNA (red: H2A.X staining -> much stronger in treated cells which have high level of damage; blue: DNA staining)

Or do they constantly "repair" also healthy areas leading to the constant replacement of the atoms

As one approximation this would be true, e.g.: proteostasis  (a process whose failure contributes to aging).

that constitute the DNA?

Note that DNA molecules are vastly outnumbered by other molecules in a cell.
DNA is quite static (see above publication), although individual parts could possibly be repaired (see above image on H2A.X);
Also there might be a selective retainment of more "healthy" DNA strands with less mutations (the immortal strand hypothesis).

Just adding my two cents about this particular point:

Others argue that tooth enamel does not regenerate or get replaced. If that is true what makes carbon dating to the year of birth +/- months possible?

Because the principle of carbon dating is not counting atoms being replaced, but measuring the same atom decaying into a different isotope of its element, which is physically totally different. Wikipedia has more info about radioactive decay.

That's not what I said. The meaning of that sentence was changed by who edited it. Look at the original text on the edition history. I've corrected it. But thanks anyway for the clarification.

Note that in the study cited in my above answer, carbon dating primarily does not measure decay of radioactivity, but inclusion of a (radioactive) isotope that happed do be created by unusual conditions (such as atom bomb tests), and its "decay / inclusion" in DNA molecules which are known to replicate in a semi-conservative way (that essentially depends upon biology and differs from pure physics / chemistry).

@CamiloRada sorry about that, I indeed did not read the original question before it was edited. I will remember to do that from now on (I started using SE only recently).


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Does natural selection in humans work out in "best outcomes" or do e.g. subjective biases affect it?
Does natural selection in humans work out in "best outcomes" or do subjective biases affect it? That is, how much is natural selection "guided by nature" and "guided by humans"?
E.g. given "fully rational" reproducers, then would the gene pool get increasingly better and produce better and better humans over time? Or is this somehow "independent" of "reasoning"? Such as that the natural selection does not follow all kinds of rationalizations.

What do you mean by "bad traits" ?

@Shred Low-worth to community, low-skill and learn-ability, low physical fitness, unattractive physical features, sicknesses.

What do you mean by "better"? And how did you come to the conclusion that evolution produces "the best"? Evolution only produces that which is able to survive, reproduce, escape the predators (sufficiently enough) - and all of this better then those competing within the same niche (https://en.wikipedia.org/wiki/Ecological_niche). If the evolution "optimizes" something, it optimizes only those traits for which there is pressure of natural or sexual selection.I recommend you to read The Blind Watchmaker by Richard Dawkins. The book is popular, but gives the understanding of the basic principles.

@HonzaZidek I think that one could hold as an abstract ideal that evolution could be altered to lead to better, improved humans. If it doesn't do it on its own. And the reason is that better humans would be even more successful? Isn't science pretty much about improving humans? But there's the difference as to up to how much can humans affect evolution, and how much of it is beyond control by humans?

I completely do not understand the term "better improved humans". All this "better" and "improved" is based only on anyone's biased view. Evolution does not operates with these terms. Evolutionary good = able to pass the genes forward. Evolutionary better = able to pass the genes forward for more generations, able to reproduce in larger scale (and still survive). Evolution-wise, our euro-american (= science-based) civilisation is not very "good", taking into account the decreasing fertility rate (http://worldpopulationreview.com/countries/total-fertility-rate/).

@HonzaZidek So then you're sort of at the center of this question. Because I'm asking whether asking for better, improved makes sense from evolutionary perspective. And up to how much. I'm not sure if improved, better are entirely useless concepts, because humans e.g. have a strong tendency towards improvement of their environment, they build tools etc. But as to whether it's through evolution or something else. E.g. what's the relationship between what each subject decides to do and evolution?

And as for your "evolution could be altered to lead to better, improved humans", this already has a name: eugenics (https://en.wikipedia.org/wiki/Eugenics), very much popularised by Adolf Hitler (https://en.wikipedia.org/wiki/Nazi_eugenics).

@HonzaZidek I would count normal human life that involves development, progress to belong to that, not just eugenics.

That is, given "fully rational" reproducers, the gene pool would get increasingly better and produce better and better humans over time.
Environmental conditions change over time. What may once have been an advantageous trait in a human population may become deleterious in some form at a later time or different environment (i.e. by spatial migration).
Case in point: Central Africa is plagued with malaria, a disease caused by a parasite of the Plasmodium genus which infects red-blood cells (RBC). Sickle cell anemia (SCA) is a hereditary disease which can be deadly, unrelated in its cause and prognosis to the Plasmodium infection, BUT, it confers a resistance to Plasmodium infections due to the RBC of SCA-patients having an aberration in the shape of their RBC's. Here is a selective pressure caused by malaria that "promotes" the profileration of SCA in the local population. As a result, what you have is a large section of Central Africans that are Sickle Cell Anemics, whom have better survival rates than non-anemics in that environment. But if you take these anemics to northern Europe or North America, where malaria is virtually non-existent (spatial, environmental change), the anemics have a higher mortality rate than the rest of the local population.
You get what I'm saying?
Natural selection doesn't produce better and better humans over time - it produces humans that are better able to survive in that era, in that place.


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Heat shock vs electroporation
I've been transforming E. coli via heat shock in order to insert oligonucleotides (around 50 nt); however, none of my experiments have given positive results so far. I begin to question the efficiency of chemical transformation, especially for short DNA fragments. Is there such a notable difference between chemical and electro transformation?
EDIT: I'm checking for CRISPR array expansion, so my oligos have a protospacer length and structure. I'm using BL21(DE3) cells which have an IPTG-inducible T7 that I'm first transforming with a Cas1-Cas2 plasmid. Then, I induce the cells in order to express those proteins, make them competent, and transform then once again but with the oligos and an eGFP plasmid (for some sort of selection). To check for positive results, I'm amplifying the leader-repeat junction of the CRISPR array and compare its length (via electrophoresis) with a usual array that has no integrands.

How do you do your chemical transformation? Do you make the cells yourself? Do you have a positive control?

Yes, I make the cells competent myself. I do have a plasmid control for antibiotic selection which, on the other hand, is successfully transformed.

In an allusion to Damir's answer below, how are you testing positive results? A plasmid with an antibiotic resistance gene is easy to test for. But 50 nucleotide oligos won't carry a useful reporter gene.

When transforming, I insert around 30 times more oligonucleotides (as mol) than plasmid, assuming that if the plasmid has entered the cell, the probability that some oligos have also entered is higher. Indeed, that's still pretty much luck, but testing hundreds of colonies and getting 0 results makes me believe that something's wrong. By the way, I'm checking for CRISPR array expansion, so for testing positive results, I'm comparing the amplicon length (which is the leader-repeat junction) via electrophoresis and after that, I'll be doing some digestions to confirm the presence of my oligos.

How do you detect a potenially positive transfection? Short fragments might have proper teriary structures which may make them behave differently than you would expect. Also, noncircular DNA and RNA is targeted much faster for degradation in most cells if the right head or tail signals are missing.
To your question: pulsed electroporation with well prepared bacteria (state, buffer,...) is usually 5-100 x more efficient than chemically competent bacteria.

See my anser above. Regarding oligo degradation (by endonucleases for example), I should mention that before transforming, I induce the expression of the necessary proteins for the CRISR adaptation stage.


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Can different Chromobactierum strains have completley different colony morphology?
In the analysis of an unknown bacterial broth I observed a gram negative rod genus. Upon plating I observed small, dark purple and convex colonies; corresponding presumably to Chromobactierum. After then replating the purple colonies I then observed white fillamentous colonies that were gram negative and was subsequently able to make an identification that suggested the organism was either of genus aeromonas, vibrio or Chromobactierum. I found an article showing that Chromobactierum colonies dont have to be purple and can have a range of morphologies (see link). My question is: upon replating, can the morphology of the two colonies change so drastically?( assuming I had two different strains of Chromobacterium present)
Source:http://jb.asm.org/content/66/4/470.full.pdf

This different Gram reaction suggests you have a contaminant.

Sorry error on my part. The purple bacteria observed on the first plate were gram negative

Did you PCR the 16S?? Most phyla are Gram negative and several skin flora bacteria are negative and unpigmented —yes, there will be many bright yellow staphs (+) and red Serratia marcensis (-), but not all. So I agree with the contamination hypothesis.


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Proof of Darwinism( fossil records)
What logic do anti-darwinists provide for refuting the proofs( by proof I mean the fossil records) of darwinism ?
Is it the missing links?
If yes then , what do they make of the fossils which are present?
Note : I'm not being hostile to the anti-darwinists, I'm just very curious.Also I really want  to see it directly from an anti-darwinist (i.e. answer from an anti darwinist highly preferred)
If there's any problem in my question please inform me.

A terminological comment; Darwinism is a rather obsolete term, if you are referring to modern evolutionary theory. Sometimes it is used in contrast to the Modern synthesis or modern evolutionary theory. Especially by creationists, and in some popular use, it is seen as a general term for evolutionary theory though. However, it is rather inaccurate to label that as Darwinism, since much of modern evolutionary theory was completely unknown to Darwin (DNA, mechanisms of inheritance, Modern synthesis, much of EcoEvo etc).

Simply calling it Evolution or Evolutionary theory is better, when you are referring to the research field as a whole.

I would also downplay "theory". We never talk about the "theory of gravity". Evolution is the only series of facts that we insist on linking with "theory", which is misunderstood by non-scientists as some kind of guess.

@fileunderwater actually I meant darwinism specifically, not the modern theory of evolutions, like the neo- lamarckian model of evolution, symbiotic model of evolution, mendelian evolution etc.

@fileunderwater darwinism is actually pretty controversial , watch this----------https://www.youtube.com/watch?v=riqCx84rhfY

@KarlKjer This is a fundamental problem. Scientists have a complete different meaning for the word theory. For them a theory a system of scientifically proven statements. For the "lay public" it is seen as an unproven hypothesis.

@user545735 Darwinism is not controversial at all. Only people who don't understand evolution think it is.

Also, if you link people to an hour long lecture on a topic, they probably don't have time to listen to it all. It's better to summarise your points in writing.

I'm voting to close this question as off-topic because, on a scientific websites, one cannot list nonsense, undefined "arguments" from creationist philosophy. If you have a specific, well defined creationist argument, we can debunk it for you but we cannot safely list and explain what is nonsense. Unfortunately, the website nonesense.SE does not exist for you to ask your question.

Related: Is evolution a fact?

@user545735 If you are asking specifically about the historical theory Darwinism, you first need to define exactly what you mean by that, and how it differs from modern concepts of evolution. Everything expressed in The Origin of Species, 6ed? The question then also becomes an exercise in the history of science.

The word "Darwinism" is antithetical to science, because it implies a system of beliefs ("-ism"). Science is not a system of beliefs, but a method to test hypotheses and data. Scientists don't "believe" in evolution or climate change. They simply find that all the data are consistent with these explanations (theories are explanations that fit the observations and cannot be falsified).

I'm voting to close this question as off-topic because it is not a question about Biology, and hence off-topic here.

I'll try to approach this topic academically and undogmatically since skepticism is essential to scientific discourse. However, bear in mind that as of $2009$, $97\%$ of scientists believed in evolution (Pew Research Center, 2009). Although I can't find how many scientists were surveyed, nor what disciplines they studied.
I've not given many citations, since I'm questioning the counter-arguments to evolution, and not necessarily providing my own specific claims. I'll also avoid discussions in comments since this could be a contentious topic.
1. Missing Links
In theory, if there aren't fossil records forming a complete chain of species from early hominids to humans, then this could cast doubt on the theory of evolution. But the fossil records are convincing and there are many of them (Wikipedia, n.d.). Also, if there truly is a missing link and there is no relation between humans and early hominids, then why do their skeletons look so similar? And why did humans suddenly come into existence? Is there a better rational or empirical explanation?
2. Skepticism of Existence of Fossils
One could propose that there are no fossil records, or that they have been faked. But why would anyone want to make any fake fossils or lie about them? What possible conflict of interest would they have, and why would this benefit them? Why would tens of thousands of scholars want to support this theory? How does this explain how scientists are still discovering new fossils? If anything, a paper that shows that evolution is false would be groundbreaking and give instant publicity to whoever publishes it.
3. Inaccuracy of Techniques: Radiocarbon Dating
A variety of scientific techniques have been used to garner information about the fossil records. One such technique is radiocarbon dating, which tells us how old the fossils are. If this technique is inaccurate, we could be wrong about the fossils and they might not form a chain that leads from early hominids to humans. But the technique is broadly accepted. Also, how would this explain the fossils - why would humanoid fossils with such odd shapes occur at random points in time? The shapes of these fossils don't look like typical human mutations.
4. Skepticism of Age of the Earth
Another counter-argument proposed to evolution is that the age of the fossils doesn't allow for their beliefs of the age of the earth. But the age of the earth is accepted by the geological community to be ca. $5\;\mathrm{bn}$ years old (Wikipedia, n.d.).
5. Summary and Micro/Macroevolution
A related counter-argument to evolution that I'd like to cover, but isn't directly related to the accuracy of fossil records is that of microevolution vs. macroevolution. Some people argue that we can only see evolution on minute scales, and not on large scales. But then if a population does have small-scale changes, and there is a higher probability individuals with certain adaptations surviving, then wouldn't these micro-effects sum, over millions of years, to produce large-scale effects?
In my opinion, the counter-arguments for the fossil records of the fossil records for evolution are rather flimsy, which suggests that evolution is the best explanation we currently have for the evidence we've observed.


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Could a human become infected with rabies in such a way that even prior vaccination wouldn't stop the infection?
Given the highly persistent nature of rabies after entering the brain, I was wondering whether certain ways of exposure to this virus could be risky even for a previously vaccinated individual. I understand that no vaccine is 100% effective, but would the risk of infection be particularly increased in case of exposure at some particular site on the body? Which mechanisms of the rabies infection would play role in this situation?

It would be helpful to further explain the reasons for the downvote so that I could improve my question or avoid asking unsuitable questions in the future (and possibly delete this one). While I welcome constructive criticism, downvote without any explanation doesn't really help anything.

There is one down vote and one close vote. I expect they're from the same user. The close vote reason is "primarily opinion based", which is probably because your question comes from a work of fiction. On Biology.SE, when you frame your question with, "I was watching this TV show... could this really happen?", you prime readers to expect your question to require speculation.  There is a real and answerable question here, though. I considered editing it to take out the TV show frame for you, but I'll let you decide if you want to do that or not :)

I don't want to put words into your mouth here, but as it stands the title text doesn't quite ask the same question as the body text - could you edit one or the other? I'd suggest keeping the question in the body.

Reframing your question as:

Are rabies exposures at specific sites on the body riskier than other sites, even for a vaccinated individual?

Possibly for post exposure prophylaxis, but probably not for pre exposure vaccination.
Rabies transmitted by a bite or scratch (almost all cases) involves a period of slow replication in muscle tissue at the site of the bite or scratch (see Murray Medical Microbiology Ch. 60 for a general discussion). This is called the incubation phase, and it is highly variable. The incubation phase is followed by a prodrome phase (virus binds receptors of sensory neurons at the site of exposure, and travels to the CNS) and neurological phase (infection of the brain).
An effective immune response is believed to be the result of both an antibody response and cell mediated immunity during the prodrome phase. Post exposure vaccination is effective because the incubation period provides enough time for a naive immune system to mount a protective response (same reference). Regardless, rabies post exposure prophylaxis (PEP) includes both active and passive immunization, i.e., vaccine and human rabies immune globulin.
Both post and pre-exposure prophylaxis failures have been reported. As discussed here cases with pre-exposure vaccination failure involve immunocompromised individuals. Cases with post-exposure prophylaxis tend to involve failure to fully follow guidelines (some are summarized here), but there is a recent case where guidelines were followed. These cases tend to involve bites or scratches to the head and face, which tend to involve a more rapid progression from incubation through prodrome to the neurological phase.
So, since reports of post exposure prophylaxis failure tend to involve specific locations of exposure (head and face), it's reasonable to assume that an exposure there has a higher risk. There is no data supporting this risk for cases with pre-exposure vaccination.

What a vaccine does is simply providing the body an attenuated (avirulent) form of the pathogen, in this case Rabies. What our immune system does is recognise the particular foreign substance and produce immune cells against it, of which some cells remain as memory cells which would trigger an immune response much faster if the same pathogen enters the host again (assuming that the vaccination was successfully performed).
However, as you know that many virus mutate very quickly and if the extent of mutation leads to production of new rabies virus which is highly distinct from the form that was used as vaccine then the host will not be able to identify the new virus as rabies.
In that way the infection may take place


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Is it true that bees and wasps don't sting if struck by hand while they are in flight?
Is it true that wasps don't sting if struck by hand while flying? I know of one person who claims to have done this at least 20 times and has never been stung. And out of curiosity (not that I'd want to kill them), what about bees?

The other answers are fine and good, but I can confirm, using solid anecdotal evidence, that it is indeed possible to be stung when swatting bees and wasps.  I've done it, and I don't recommend it.

I agree with @Amory here as I know of folks being stung when driving cabrio cars. Not struck by hand but by a 80 km/h headbutt :)

I do this often and never got stung, but I saw someone else get stung by a wasp when waving it away from their face - even though the wasp didn't land on their hand at all. So it's mostly matter of chance and speed at which the hand moves. I successfully killed many wasps by clapping my hands on them in flight. But when some got between the fingers, I got stung.

I can confirm that I have often enough slapped a wasp or bee in flight without being stung. The reason is probably pure chance.
It is kind of like asking if a human can bite when slapped. If I come up to you and slap you, fast and hard, you will probably not be able to bite my hand because of surprise and the blinding speed with which I hit you. If, however, you happen to be talking to me and have your mouth open at the time, you might be able to pull off a quick bite.
In a similar vein, if I happen to swap a wasp (no anagram intended) and hit it on the sting, I will probably be stung. If I hit it on the side and move fast enough, I doubt it will have the time to reorient itself and sting me before my hand is gone.

I have many experiences with bees. I avoid swatting bees/wasps, since if you do not agitate them, they fly away and are not interested in humans or exploring human skin. (They are also important pollinators.) If you slap them very hard, you can instantly kill a bee, especially if it collides perpendicularly with a wall afterwards. If you swat it a bit less vigorously, like I did yesterday, my finger got slightly red and itchy; I was partially stung and/or the venom was transferred to my finger, which is normally stored in a cuticular sac in the abdomen of the bee and is quite potent, given its small volume.
Don't hurt bees and they can't hurt you. If you agitate bees, they will defend themselves. Any insect of sufficient size and mass can be hurt with a blow of sufficient energy. Tiny fruit flies are too small for a slap, large moths, bees, dragonflies can be slapped down when in flight.


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Injuries inflicted to Tyrannosaurus by Triceratops
I encountered this YouTube video:
YouTube video by ExtinctZoo
The video presents Triceratops as the only dinosaur documented to injure a Tyrannosaurus (excluding other T. rex individuals). Specifically, the following section mentions two fossils of Tyrannosaurus with signs of injuries probably inflicted by Triceratops.

The Dueling Dinosaurs: the famous fossil shows a juvenile T. rex with jaw injuries among others next to a Triceratops with a bite mark. This probably indicates that the two fought with the Triceratops inflicting more damage. It is unknown what has killed both.
A T. rex skeleton nicknamed "rex" (?) of a 12-meter individual with a puncture injury in its femur, where the hole matches a ceratopsid horn, with the major candidates are Triceratops, Torosaurus, Ojoceratops with Triceratops being the most likely one (because it was the most abundant large dinosaur of its habitat).

However, I could not find paper references to these fossils with the description of the T. rex injuries and the association of them to Triceratops' attacks. Do you know on which papers the movie maker is based and if there are, in addition to the full texts, open access or articles in popular science online magazines which I can refer to and indeed assert the injuries to Triceratops attacks.

I will start with the dueling dinosaurs specimen in which a Nanotyrannus lancensis (BHI 6437) and unnamed Chasmosaurine ceratopsian  were found together with good evidence for dying together. Not a triceratops but early description was of a triceratops so the confusion is reasonable.
The specimen has a strange and legally problematic history going through both private and public hands so published study has been non-existant. The hope is with their new public home more work will be done. Scientists don't like to publish on private specimens since access is not guaranteed so many publishers will not accept such work. On top of that a lot of paleontology publications wait until the yearly Society of Vertebrate Paleontology convention each year; with the grand opening in April of 2024, I do not expect much publication until the fall.
Hopefully it is real specimen, possible fakes is one the big risks with private collections and another reason it is strongly discouraged. If it is real it will represent a truly amazing find, but with no publication, ascertaining the truth of the injuries is impossible.
the lack of publication is also the reason for the lack of pictures "poaching"  publication occasionally happens so don't expect many detailed images until it gets published. This can even happen accidentally with new species because the first name given in publication is the one that sticks. So it is common for researchers to be protective of diagnostic images until the first publication.
https://www.bonhams.com/auction/21076/lot/1032/dueling-dinosaurs/
https://web.archive.org/web/20201117124307/https://www.nationalgeographic.com/science/2020/11/dueling-dinosaurs-fossil-finally-set-to-reveal-secrets/
current location https://naturalsciences.org/exhibits/permanent-exhibits/dueling-dinosaurs
I know of no T-rex specimen named "rex" nor of one with a punctured femur, nor could I find any publication of either. So it is either not a T-rex or is not published yet.

Thank you for your answer. I'll wait someone with more information will comment.


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Why Triceratops' frill reabsorbed the triangular epiossifications on the edge of the frill?
Why Triceratops' frill reabsorbed the triangular epiossifications on the edge of the frill?
Juvenile and even sub-adult Triceratops had triangular prongs on the edge of the frill, called "epocciptials" or "epiossification". However, in very mature specimen the frill lacks those triangular prongs and transitional specimens show how the epiossification fuse with the frill to form a wave-like edge and later even a smooth edge.
A scientific description of this ontogeny can be found here:

Horner, John R; Goodwin (2006). "Major cranial changes during Triceratops ontogeny". Proceedings of the Royal Society B (273): 2757–2761. doi:10.1098/rspb.2006.3643.

Short summary by Horner: https://www.youtube.com/watch?v=m2_Fcp6W7U4
The question is: why? What benefit did this modification (the smoothening of the frill's edge) give to Triceratops?

General Images

Figure 1.1: this Triceratops exhibits prominent triangular epiossifications. Source: https://en.wikipedia.org/wiki/File:Triceratops_skull_frills.jpg

Figure 1.2: growth serie of Triceratops.
Source: https://en.wikipedia.org/wiki/File:Triceratops_ontogeny.jpg

Figure 1.3: a skeleton of adult Triceratops in New York's AMNH. The frill shows a smooth edge.
Source: https://en.wikipedia.org/wiki/File:Triceratops_AMNH_01.jpg

Triceratops horridus

Figure 2.1: Triceratops horridus Marsh, 1889, shows scalloped edge https://www.si.edu/object/triceratops-horridus-marsh-1889%3Anmnhpaleobiology_3383580
Figure 2.2: https://commons.wikimedia.org/wiki/File:Triceratops_horridus_skull_in_the_Gallery_of_Paleontology_-_Paris.jpg
Shows transition from scalloped to wavy edge.

Figure 2.3: Triceratops horridus showing wavy-to-smooth edge.
https://en.wikipedia.org/wiki/File:Triceratops_Hendrickx2.jpg

Figure 2.4: a skeleton of adult Triceratops in New York's AMNH. The frill shows a smooth edge.
Source: https://en.wikipedia.org/wiki/File:Triceratops_AMNH_01.jpg

Triceratops prorsus

Got a couple of different species there. Bottom photo is Triceratops horridus; top two photos are T. prorsus I think, though could be other ceratopsids. Fusion of bones is common with age - our skulls do it.

As Bob out the bottom image is a different species. So the comparison is faulty.
as for why bones fuse with age, that is very normal, it happens to bone in thousands of species our own included. The parts are separated by growth surfaces, when growth slows down aka adulthood, those surfaces fuse for strength. This is so predictable in humans the fusion of different bones can be used to age a skeleton. We see this in dinosaurs too but without living one to correlate fusion times with it can't be used to age specimens well.
As for why the angle of the epocciptials change , that's easy, the number of epocciptials do not change so as the frill gets larger the epocciptials need to get wider. They can either grow in all dimensions or just get wider making the angle shallower. Whatever function the epocciptials have must not be impacted by getting shallower so that's what they do saving bone and horn. considering how much mass they would add if they grew in all dimensions its not even that surprising.
https://www.sciencedirect.com/science/article/abs/pii/0305440388900829
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6427487/
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3864909/

That bones fuse with age, I know. The question is why the triangular epiossifications reabsorbed and disappear and not merely fuse but retain their shape. The scalloped frill of a Triceratops has a defensive advantage and probably also a display value from the prominent epiossifications in other ceratopsids).

They don't, you are looking at two different species. if you count the number of epocciptials they stay the same in the same species throughout its life. in Horridus the epocciptials are tiny, so they are just really hard to see in the adults as their width is drastically greater than their length.

I added Figs 2.1-2.4 consisting only of Triceratops horridus (however, I could not determine from the images what was the age of the animal it came from).

@Triceratops from my skimming of the literature it seems you are not alone here - the problem is that formerly there were a bunch of species, all classified on the frill edges. However, with the recent clumping, there is now diversity in the frill edges within each species - so some have it and some don't within the same species, this means that it likely isn't age dependent, so your premise fails to some extent there.

@bob1 if there is a variation within a species it is likely sexual dimorphism. but if you look at the series in fig 2 the epocciptials do not disappear. the only figuire without epocciptials is 2.4 which could be not collected, a diffrent species, or even destroyed it is a very old specimen. If you think they disappear you will need to demonstrate that.

@John I'm no expert on dinosaur systematics at all, so I don't know that they disappear or not, just that there is some diversity in size/shape across a species. Sexual dimorphism is definitely a possibility, that hadn't occurred to me.


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How do whales swallow food underwater without drowning?
How do whales swallow food underwater without drowning?
I know they have a valve that blocks the trachea while the mouth is open and that breathing is made from dorsal or parietal breathing holes or nares, which have valves that close underwater. But when they eat underwater, how do they prevent seawater from flooding their stomach and drowning them?
Crocodylians face similar problems. They have a palatal valve that seals the trachea and the esophagus while it opens their mouth underwater, preventing them from drowning. However, when feeding, crocs swallow above water, putting their head out of the water. That's how they prevent water from flooding their stomach.
I reckon that whales who eat fish and squids do not resort to this solution, since it will require them to surface after each fish is swallowed. However, Baleen whales filter the water out using the baleens in their mouth, but what about "teeth" whales like orcas?

Your title and question are both poorly worded. You alternate between talking about drowning and swallowing large amounts of water and treats the two as if they were the same thing. They are obviously not. Since it is well within human experience to swallow things without also choking on them, I think you are more interested in how whales do not swallow a bunch of water, and if they do, how they deal with it.

You seem to have Googled about baleen whales and crocodiles so I assume you tried Googling about orcas too but were having trouble. Trying Googling about "dolphins" instead. The answer comes up right away.

@DKNguyen - you are borderline answering the Q in the comments; I would suggest turning it in an answer, because it already is an interesting read as is :)