text:
What is the current scientific consensus regarding the relationship between Eukaryotes and Archaea?
In the traditional 3-domain system, the domains Bacteria, Archaea, and Eukaryota are all distinct from one another, with the latter 2 usually being sister-groups in a clade.
However recent evidence has come to light showing that Eukaryotes might have arose within Archaea, rather than being the sister group to it. More specifically, Eukaryotes might have evolved from the Asgard archaean superphylum.
What is the current consensus towards this issue? Do most evolutionary scientists today still consider Eukaryotes to be the sister group to Archea? Or is it thought that eukaryotes are archaeans?

What makes you think that there is a consensus on any controversy in science, and what makes you think that knowing which view is dominant has any importance per se. As scientists, we on this list are only interested in the strengths and weaknesses of arguments. Please present your questions in these terms.

For a recent provocative article on this subject, see Isolation of an archaeon at the prokaryote-eukaryote interface, and see also The mysterious microbes that gave rise to complex life (both articles published in Nature in 2021)


text:
In evolutionary history, which came first - neuron or astrocyte?
As research is showing astrocytes role in information processing [1], I am interested in knowing which evolved first - an astrocye or a neuron ?
[1] Santello, M., Toni, N. & Volterra, A. Astrocyte function from information processing to cognition and cognitive impairment. Nat Neurosci 22, 154–166 (2019)

Short answer
Neuroglia evolved after neurons did. The evolution of glial cells coincided with increased complexity of the organization of the nervous system.
Background
Glial cells are supportive cells for neural tissue. As such, evolution of glia before neurons would not be a parsimonious hypothesis, however not impossible.
Glia indeed seems to have evolved after neurons did. Arguably, Ctenophora (comb jellyfish) and Cnidarians (hydras and sea jellyfish) feature the most primitive neural systems known today. They have diffuse nervous systems, characterized by relatively simple semi-independent networks. These networks lack conglomerates of specialized neurons called ganglia, and these animals don't have a brain or a spine such as the chordata have. They also don't have glial cells associated with the diffuse neural networks.
Only until more complex nervous systems appeared later in evolution, i.e., in early Bilateralia (e.g., flatworms and roundworms) supportive cells evolved: the neuroglia.
Reference
- Verkhratsky et al. Adv Exp Med Biol (2019); 1175:15–44

Fig. 1. Evolution of glia accompanies increased complexities of early life forms. source: Verkhratsky et al. (2019)

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text:
Does Telomere length shortening with age actually cause our cells to age and stop functioning properly?
The human telomere, a simple repeating sequence of six bases, TTAGGG located at the ends of chromosomes (Moyzis et al, 1988) protect them from degeneration, reconstruction, fusion, and loss. It is believed that  this shortening becomes critical for a telomere on a particular chromosome, which becomes unstable and the cell stops dividing.
Nevertheless, as human dermal fibroblasts of 100-years-old individuals still have a telomere length of 6–7 kb and retain proliferative capacity for about 20 doublings (Allsopp et al, 1992). They have not reached the limit of their proliferative capacity or of telomere shortening even at such an advanced age. The telomere length of human peripheral blood lymphocytes decreases with age and shortens to about 5 kb in some individuals after age 60 years, but no individuals have been found with a mean telomere length of less than 5 kb, even among 100-years-olds. (Vaziri et al, 1993) (Iwama et al,1998)
Does this Telomere length shortening actually cause our cells to age and malfunction?

Yes

"all somatic cells have a molecular clock that leads them to develop a senescence phenotype" Ruiz et al. 2022

Short telomeres cause replicative senescence. Senescent cells upregulate survival factors and do not undergo apoptosis (although they should). Senescence is accompanied by SASP (senescence-associated secretory phenotpye). A cell with SASP will secrete inflamatory (and cytotoxic) factors that attract immune cells, maybe in order to be killed and removed from the tissue. If senescent cells aren't removed (due to a high burden of senescent cells), these inflamatory factors also damage surrounding cells. This accumulation of tissue damage is pretty much what "aging" is.
However, there are other mechanisms that cause senescence/aging: There is also DNA-damage, mitochondrial DNA-damage (causes production of reactive oxygen species), accumulation of protein aggregates, chronic inflammation, etc.
Note that cellular stress (drug therapy, genotoxic stress, irradiation, inflammatory stress, etc.) can accelerate telomere shortening. For example, smoking during pregnancy  reduces telomere length of the unborn child (Mirzakhani et al. 2017)
But still, telomere length is only a  "rough estimate of aging rate and can hardly be regarded as a clinically important risk marker for age-related pathologies and mortality" (Vaiserman & Krasnienkov 2021)
Further reading: Chaib et al. 2022

Yes.
This was demonstrated by showing that re-activation of telomerase (which lengthens telomeres) prevents cell senescence.
The immortal nature of cancer cells can be attributed in large part to their constitutive expression of telomerase.
The mechanisms by which short telomeres lead to / are linked to senescence, aneuploidy, etc.  occurs have been pretty well worked out.

Yes, it's true in eukaryotes after every cell division telomeres shorten by many base pairs. Telomerase is not found in prokaryotes as their chromosomes are circular. So, no DNA is left out for copying.
In eukaryotes, telomerase is found in stem cells.
Note: There are different types of stem cells that exist in humans at every point of their lifetime. But none of them are totipotent(those have the ability to create a full organism)
Thus, telomerase ensures the safety of genetic information by elongating the telomeres.
Mechanism of telomere regeneration given in AP biology...I m posting the link below
https://openstax.org/books/biology-2e/pages/14-5-dna-replication-in-eukaryotes

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What do the colors mean in representations of amino acids?
I've seen sequences of amino acids depicted using colors, which I imagine correspond to whether the side chain is charged, etc. I can't figure this out, though, and I'm not a biologist or biochemist. I've seen what appear to be some different color schemes online. Is there a standard format / scheme such as for the following example?

I don't think there is a general standard for the color code of the amino acid – there are different conventions and software or paper authors may decide to use their own. You can see some examples of color codes here. As for your particular example, I would say it is more or less a custom color code, made for this particular case.

There is no ‘standard’ colour scheme for amino acids in the sense of one recommended by a standards or professional organization for biochemists. There are several schemes used in practice — either by software applications or individuals — based on two main criteria:

The properties of the amino acid the colours are intended to indicate
The colour choice of the individual author in expressing the intended similarity

The graphic below is taken from a page I found on the web by a random search and relates to a particular software package. It illustrates some possible colour schemes, none of which correspond to that in the question.

The ‘Chemistry‘ scheme focuses on one view of chemistry — acidity, basicity, aliphaticity, aromaticity, polarity (amides etc.) although it has had to make a choice in classifying tyrosine with tryptophan rather than threonine. The ‘shapely‘ scheme focuses solely on size and shape, whereas the hydrophobicity scheme does as its name implies. The ‘Zappo‘ and ‘Taylor‘ schemes (never heard of them previously) seem to be individual slight variants on the first two. The letter scheme appears to be just a way of distinguishing each amino acid for identification purposes, and molecular graphics software (e.g. Jmol) generally lets one colour amino acids along a protein in a graduated manner from (e.g.) red to blue to indicate the pathway of the chain (e.g. in this previous question on the list).
Oh, and I adopted another compromise scheme (probably using colours easily available in Excel) for a paper I published in 2021:

Coda
There is a tendency in schemes that reflect the chemistry of the amino acids to use blues for basic amino acids and reds for acidic ones. This possibly reflects the widely used chemical CPK colour code of blue for nitrogen (as in amino groups) and red for oxygen (as in carboxyl groups), and might have been regarded as an aid to recognition. (A simpler explanation might be a relationship to litmus paper.)


text:
What is the physical dimension of international unit (IU)?
I was skimming through a study [mRNA-based COVID-19 vaccine boosters induce neutralizing immunity against SARS-CoV-2 Omicron variant, page 6, figure 3(A)] on vaccine and I came across the international unit ($\text{IU}$). From what I read, $\text{IU}$ measures the amount of substance adjusted according to biological activity observed. [I studied maths in uni. I only learnt a bit of physics/chemistry/biology in secondary school.]
In dimensional analysis, all units are assigned a physical dimension which is expressed in term of length ($\text{L}$), mass ($\text{M}$) and time ($\text{T}$). For examples, gram has dimension $[\text{g}] = \text{M}$, mole is dimensionless, millilitre has dimension $[\text{mL}] = \text{L}^3$ and molar has dimension $[\text{M}] = \text{L}^{-3}$.
What is the physical dimension of $\text{IU}$?

@ARogueAnt. The $y$-axis of the graph has the label Pseudovirus neutralization (IU/mL). I don't know that it varies according to the substance measured. It's something new to me.

Please link the study

@acvill mRNA-based COVID-19 vaccine boosters induce neutralizing immunity against SARS-CoV-2 Omicron variant, page 6, figure 3(A)

It depends totally on the substance. IUs are decided upon by committee in order to be able to compare formulations.

As you have identified it yourself, the $\rm IU$ is a measure for the "effective" amount of substance, which is one of the seven base quantities in the International System of Quantities. Therefore, in your notation, $[{\rm IU}] = {\mathsf N}$.

Based on the WHO mandates, the international units ($\text{IU}$) are arbitrary units corresponding to neutralization. The reason for choosing arbitrary units was that the serum used to test immune resistance is made of too many molecules for there to be a single standardized piece to measure. As a result, the units are arbitrary and that means it is based on the number of particles neutralized, so the $\text{IU}$ has a dimension of $\text{[IU]}$ = $\text{N}$, $\text{N}$ being the number of neutralized particles.


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What insect/invertebrate species evolves fastest?
I am starting an experiment in which I will be forcing evolution in a moderately complex species of insect or invertebrate. I am prepared for the possible longevity of this experiment, but i have no clue what species i should use. I would prefer a species that can easily respond to biotic challengers. Reason being, every abiotic factor will be promoting the growth and prosperity of the target species, however every biotic factor will be trying to kill the target species. The species can be aquatic, amphibious or terrestrial.

Drosophila melanogaster. (fruit-fly, pomice-fly)

Image, public domain, via Wikipedia 2022.
Development time is under ideal conditions 8.5 days (at 25 Celsius, 77 Fahrenheit), the females produce perhaps 500 eggs per generation which can hatch in 12-15 hours - from the point of view of turnover, you should be able to observe many generations in a short-ish period at one generation per ten days in reasonable conditions.
Their diet is fruit, fungi - both preferably decomposing and are able to be anaesthetised with ether or carbon dioxide.
They have acted as a model organism (eukaryote) for the study of genetics and there are many known mutations, and established ways to produce them (ethyl methane sulfonate, ionising radiation).

A classic example

Thank you very much, you gave me all the info I needed.

@Astrophysicsteen You're welcome. Feel free to "click the tick" if you find this appropriate as an accepted answer.


text:
Is there an established algorithm to produce a hash number from a person’s DNA?
It is often said that a person’s DNA uniquely identifies that person. Is there an established algorithm that can/could take values of a specific DNA sample to produce a digit string unique to its source, perhaps as a hash of the quantification of DNA components?
I am a software engineer with very little knowledge of genetics, so I’ll stipulate this may be a dumb question for those skilled in the art.
However, as hash spaces are easily defined that can uniquely represent trillions of elements, it seems me that this should be possible and likely has been worked out.

This rather assumes 100% copy fidelity in every cell of the body as a basis, that's not quite the case.

One slight correction. Not all humans have different DNA sequences — identical twins start out the same in this respect.

As a link to a paper is not an answer, and I cam not competent to summarize the paper, I just provide this info in a comment. Have you seen Zobrist's "Ordered Minimal Perfect Hash of the Human Genome and Implications for Duplicate Finding"? It requires membership of IEEE or institutional access.

There are various approaches based on sequence k-mers that do more or less this. I don't off the top of my head know which ones are the most trusted for minimizing collisions, which appears to be your motivation. For a general purpose k-mer library you might look at khmer. For something a little less biology focused you might look at murmurhash, which I believe is nonetheless used extensively in bio applications.
MinHash has been used fairly extensively to generate sequence hashes efficiently in large nucleotide sequence collections. However, it relies upon a summary that may not guarantee uniqueness at the level you are interested in.
For a thread discussing different approaches and with more background, see here.
If you are interested in human sequences specifically, there is a good amount of material out there on constructing unique hashes, though I'm not personally so familiar with it. Here is one reference at google-random, here is another.
It's also true as stated in the comments that the sequence of an individual is noisy. I would argue that there is a strong enough central tendency in the distribution of sequences for a human that it's still a worthwhile exercise, but understanding this shortcoming will be crucial for interpreting data (especially given the error-prone nature of sequence reconstruction via sequencing and assembly).


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What is a (preferably) flying carnivorous insect that hunts adult fruit flies at a rate that doesn't threaten extinction?
I am forming an experiment, where I will be forcing evolution on Fruit Flies
(Thanks rogue ant) The experiment goes a bit like this;
I will be forming an ecosystem where every abiotic factor is perfect for the fruit fly, and is supporting its growth and species prosperity. everything down to temperature and humidity, however every biotic factor is trying to kill the fruit flies.
If anybody knows of a species that would work for this, or has miscellaneous tips, or ideas, please let me know.

You could also test whether the lab strain of flies have lost any predator avoidance behaviours compared with wild-sourced flies.

Good idea! Will surely look for changed behaviour patterns.

Biotic fruit fly pathogens / predators.
You don't need every single one for a basic project.  Here are 2 types.
Parasitoid wasps
Parasitoid wasps lay eggs in larva.

Drosophila in the wild suffers massive mortality from the attacks of
parasitoid wasps. As many as 80% of Drosophila larvae in natural
environments may be killed by wasps that lay eggs in them. These eggs
hatch into larval wasps that devour the fly larva from within. Adult
wasps then emerge and mate, after which female wasps search for new
hosts in which to begin the cycle anew.

source
Earwigs, spiders and ants oh my!
Predation on Drosophila suzukii within Hedges in the
Agricultural Landscape

When we exposed D. suzukii pupae in dry and humid hedges, we found
that on average 44% of them were predated within four days. The most
common predators in the hedges were earwigs, spiders, and ants.

There are lots of other mortality causes for fruit flies - fungal infections, bacterial infections, mites, viruses.  But as far as predation these will do.  The parasitoids for the larva and the 3 predators for adults.
Plus those 4 insects will make a sweet icon.

While I appreciate the contribution, I was mainly looking for something that preys on the adult flies. I will edit the post to mention this, Thank you though!


text:
vitamin D in mushrooms
How could I test vitamin D levels in mushrooms? (I am a high school student, our lab is pretty modern, but my resources are limited)
I found an article about it (https://www.hindawi.com/journals/jfq/2020/8869279/) but I cannot understand it. Also, it is on a few different types of vitamin D. I want to test the active(usable?) vitamin D amount in different grown mushrooms.

Welcome to Biology.SE. This is not simply an answer site, but instead a site that promotes self-learning with some expert help. Consequently, questions that show little or no prior research effort are off-topic on this site as are "homework" questions unless you have shown your attempt at an answer. Please [edit] your question and tell us where you've looked for answers, what you do know about the topic, and where exactly you still have questions. Please take the [tour] and consult the [help] starting with [ask] for details.

In addition, we have no idea what "pretty modern" means — if you want useful answers you need to put in more effort on your part. Do you have access to an HPLC/FPLC machine or any kind of set up for doing column chromatography? In not, then the paper you linked is of no help to you. Have you researched what forms of vitamin D are "active"? If not, that would seem to be your first step — I suggest editing that information into your question along with what you can find on your own about methods for detecting those compounds.


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Would it be possible to genetically modify a plant to produce usable antibodies? And if eaten would they be used?
I am thinking of a project where i would Gene edit a plants cells to produce anti bodies that humans could use. Does anyone know
A. what genes to modify?
B. Would the antibodies be usable after ingestion?
C. What plants should be used?
(I know next to nothing about botany)

Possible duplicate https://biology.stackexchange.com/q/56131/27148

Welcome to Biology.SE. This is not simply an answer site, but instead a site that promotes self-learning with some expert help. Consequently, questions that show little or no prior research effort are off-topic on this site. In addition, questions must be focused — you have three distinct questions. Please [edit] down to a single question, tell us where you've looked for answers (including links to related questions on this site), what you do know about the topic, and where exactly you still have questions. Please take the [tour] and consult the [help] starting with [ask] for details.

To answer the "plants producing antibodies" part:
Yes.

in 1989 scientists developed a technique to genetically alter a
tobacco plant so it would produce monoclonal antibodies

Researchers had to adjust the West Nile virus antibody to adapt it for production in Nicotiana benthamiana, a relative of tobacco:

"We altered the genetic coding of the antibody slightly, not changing
its parts but using alternate forms of the coding for those parts to
maximize the plant's ability to produce it," says co-senior author
Qiang Chen, Ph.D., of Arizona State. "We also stabilized the antibody,
preventing copies of it from being degraded inside the plant cells.
Together, those two techniques increased our average antibody yield by
60 percent above any previous efforts."

Although, the study acted as "proof of concept", the antibodies in question were only a partial success:

They found that one important receptor was binding appropriately, but
the strength of a second receptor's bond was lowered.
"This results from the fact that plants combine their proteins with
slightly different sugars than mammals," says co-senior author Michael
Diamond, M.D., Ph.D., professor of medicine, of molecular microbiology
and of pathology and immunology at Washington University. "We're
already working on genetically modifying the plants to humanize the
sugars the plants combine with the antibody's proteins."

The (successful) release and usage of ZMapp in 2014 against West-African Ebola virus outbreak virus furthered the public awareness of the technology.


text:
Cloning in a large DNA fragment into a plasmid
I need to clone in a 30kb DNA fragment into my plasmid (~7kb).
Working with such a large fragment I have run into  a few problems. The first was purifying it from the PCR mix used to amplify it out of its template, I achieved thsi with ethanol precipitation however I am unsure if thsi removes dNTPs from the PCR mix?
The second is ligating it into my plasmid. The fragment is flanked with EagI sites, however after digestion I cannot purify the digested fragment from the end cut fragments(due to its size).
I have two questions, firstly will the dNTPs remaining from the ethanol precipitation affect digestion and ligation reactions. Secondly will the end cut fragments affect ligation reactions.

Is your plasmid capable of handling such a huge insert? Most of the commonly used plasmids are not.

Welcome to the site. Can you please edit in some more information (the more the better for troubleshooting) - how big are your fragments from the EagI site digestion? How are you trying to separate the digested from undigested? Have you looked at things like Bacterial artificial chromosomes (BAC) that might be more appropriate for this size. What is the purpose of the cloning - expression, (cloning) library prep...?

Hello,
Yeah so my plasmid is a bit of a weird guy, its linear (has covalently closed ends) and has been demonstrated to be able to hold up to 40kb without too much problems. I've also further modified it to potentially make it hold more.
So the plasmid (being linear) has a left arm that it 5.7kb and a right arm that is 1.1kb; these are treated with EagI to leave overhangs. The fragments I am trying to clone in is 30kb and has eagi sites at each end.

So this plasmid is one which I have modified from a larger linear plasmid. The whole purpose of this is to be able to store and propagate large and complex DNA fragments without needing a very low copy number plasmid. The specific purpose of it is quite broad, and could be used for many purposes. Basically trying to demonstrate that my engineered plasmid can hold such large and complex fragments.
At the moment I'm assuming the fragment is being digested completely, and I am not doing a cleanup after because they don't seem to be able to cleanup such a large fragment.

The problems I'm encountering are about when I'm trying to ligate things into it, as this big 30kb fragment is hard to isolate using commercially avaliable PCR cleanup kits and so on.

Please [edit] your post to summarize the additional information you've provided in the comments. Comments are ephemeral and often overlooked so they should only be used for discussion of how to improve the associated post. I would suggest also including some information about the plasmid (with citations). Finally, you have multiple questions some of which I'm sure you can answer yourself — please edit this down to one clear question (make sure the title reflects that final question). That will help you get an answer and make this more useful to future users.

Please also take the [tour] and consult the [help] starting with [ask] for details about how this site works and what is expected of users. This will help you get good answers (by asking well-formed, appropriate questions) and mitigate some of the frustration that new users often experience on SE sites.


text:
How do tapeworms aid weightloss?
The 5 second answer people come up with is, "well they consume calories you otherwise would have". This logic works great if we're discussing the impact of my brother consuming part of my meal but it does not however in this case. The speaker seems to have forgotten that the tapeworm resides inside of you. Any calories which aren't incorporated into your tissues are instead incorporated into the tapeworms tissues. Since the tapeworm is inside of you, your weight measured externally (i.e. including the tapeworms weight) still increases.
Analogizing again to my brother: there would be no difference in my final weight gain if a) I ate dinner and then my brother or b) my brother ate my dinner and then I ate my brother : ).
So, what is the explanation for why tapeworms lead to weight decrease?

Your question presupposes that they do help with weight loss (as opposed to being a folk-tail or quackery with no evidentiary backing). They are know, among other things, to increase appetite and weight-gain in many sufferers. Can you link to any study which supports your underlying assumption?

Tapeworms are parasites that live in the gut and absorb food. They lead to weight decrease because they reduce the amount of food that can be absorbed by the body. Tapeworms also cause malabsorption, which is when nutrients from food are not properly absorbed by the body. This can lead to deficiencies in essential vitamins and minerals, which can also cause weight loss.


text:
What prevents non-aminoacylated tRNA from binding to mRNA on the ribosome and disrupting protein synthesis?
Specific aminoacyl-tRNA synthetases catalyse a reaction in which a transfer RNA molecule with a given anticodon is covalently attached to its cognate amino acid (aminoacylated).
What factors favor the binding to the ribosome/mRNA of an aminoacylated tRNA molecule, rather than one without an attached amino acid?
Is it exclusively the presence of an exposed OH group on the end of the non-aminoacylated-tRNA molecule or is it possible that the tRNA molecule undergoes some degree of conformation change when the amino acid is enzymatically attached that makes the anticodon have a greater affinity for the codon?

I made a few edits that I think make your question clearer — please check and [edit] if you feel I have altered the question you are trying to ask. Note that binding and bonding are (in biochemistry/molecular biology) used differently — bonding suggests a covalent bond was made, while binding covers other interactions. Also, please make sure you're familiar with the steps of translation — this is covered in many textbooks (e.g. search the NCBI 'bookshelf'). I think you may find clues (if not a complete answer) by doing so.

Yes I am familiar with translation.  I think my question was overly developed.  What I really want to know is what mechanisms keep a t-RNA that is not bound to an amino acid from getting into the ribosome and sticking to the codon.  I have read that the exposed OH group on the non AA bound tRNA is repelled by the ribosome, but also hypothesize that the t-RNA may have its shape altered when it's bound to an AA to make the anticodon lock up tighter.

I think the question is good now.

Great. Do you have a reason to think exclusion must be happening? The reason I suggested reading up on translation is because IIRC the amino-acylation of a tRNA creates a "high-energy" molecule, which is required for the addition of the amino-acid to the C-terminus of the growing protein. If it isn't charged I suspect (haven't had time to check) the tRNA won't have a large effect on translation ...

@tyersome I am not asking why the uncharged, non amino acylated tRNA can't contribute to translation.  I am asking why an uncharged, non amino acylated tRNA molecule doesn't occasionally slide into the ribosome and bind to a codon.  It probably doesn't matter.  It would eventually detatch.   What actual changes occur in a t-RNA molecule when it gets charged?

I posted a (brief) answer below, let me know if it helps. The first reference is fairly readable, so (depending on your background) you may find it to be of some interest. If you could post where you read that the uncharged aa is repelled, I can have a look later — my "quick" literature search could easily have missed something important.

I have edited the question in the following ways. 1. I shortened title without changing meaning in any way. 2. I inserted precise biochemical terms where needed. In particular I replaced the word "bound" in relation to amino acids and tRNA by (covalently ) attached or aminoacylated. This makes a clear distinction with the non-covalent binding of the tRNA to the ribosome. 3. I also made clear that the tRNA binds to the ribosome as well as interacting with an anticodon, as was indicated to be the concern of the poster in a comment.

Short answer
The codon-directed non-enzymic binding of tRNA (aminoacylated or not) to the A-site of the ribosome is much weaker than the (normal) binding of aminoacyl-tRNA complexed to EF-Tu/EF-1, the tRNA-binding elongation factor (which discriminates against non-aminoacylated tRNA). Hence it is unable to compete effectively with the latter to disrupt protein synthesis.
Explanation
The stages in protein biosynthesis relevant to this question are shown below.

In stage 1 tRNAs are aminoacylated with their cognate amino acid in a reaction catalysed by a specific aminoacyl-tRNA synthetase.
In stage 2 aminoacylated tRNAs (except the initiator tRNA) are recognized by a single elongation factor (EF-Tu in prokaryotes, EF-1 in eukaryotes) and form a complex with it and GTP. The elongation factor will not form a complex with non-aminoacylated (also referred to as ‘deacylated’) tRNA, and this has been shown to be caused by structural differences from aminoacylated tRNA. This latter is particularly pertinent to the question, and will be discussed in more detail below
In stage 3 this complex binds to the A-site of the ribosome in a codon-specific manner. However it is important to understand the strength of the interaction between the elongation factor and the A-site of the ribosome compared with the tRNA–anticodon interaction alone. The latter is, of course, necessary for accurate protein synthesis, but may be regarded as preventing (or  more strictly delaying) dissociation of the complex.
Basis of the discrimination against non-aminoacylated tRNA
The discrimination occurs at the stage of binding by EF-Tu/EF-1. What is known about its structural basis? This appears not to be merely recognation of the amino acid, but involves indirect effects on the structure of the tRNA and its recognition by EF-Tu, as is discussed in the 1996 paper of the Aarhus group that elucidated the structure of the aminoacyl-tRNA.EF-Tu complex. I quote:

Deacylated tRNA binds to EF-Tu-GTP with an affinity which is approximately four to five orders of magnitude lower than that of aa-tRNA. Thus, the aminoacyl group is a primary discriminator in the ternary complex formation. It is impossible to explain this by the direct interactions with the aminoacyl group alone. Other structural features of aa-tRNA must contribute to the affinity.
There has been a long-standing research on the conformational changes of tRNA upon aminoacylation. Fluorescence studies… have indicated that conformational changes occur upon aminoacylation, though in a diverse manner among the individual acceptors.

One precise structural difference is discussed:

It is evident that aminoacylation of tRNA restrains the conformational space of the terminal A76 considerably. Compared to the crystal structure of the ternary complex, the residues A73 through C75 of deacylated tRNA-Phe (PDB entry code 4TNA) are in an equivalent conformation, though slightly shifted in their position relative to the acceptor helix. However, the terminal A76 residue in deacylated tRNAPhe adopts a conformation which is impossible in the phenylalanylated form (fig 7).

[Non-acylated free tRNAPhe (left) and acylated Phe-tRNAPhe of the ternary complex (right)]
It can be seen that the residues 73–76 are near the 3′ end of the tRNA where the phenylalanine (Phe) is attached.
Evolutionary perspective
One presumes that primitive protein synthesis did not involve elongation factors. It would have been important to prevent the unproductive binding of un-aminoacylated tRNA to the ribosome and the competition with aminoacylated tRNA postulated in the question. A first step towards this may have been the evolution of a ribosomal A-site that could discriminate between the structures of aminoacylated and non-aminoacylated tRNA. The development of tRNA-binding elongation factor, as well as making the process more efficient would have amplified this discrimination which is in the range of  10-200x, depending on the concentration of magnesium ions (which artificially enhance binding of non- aminoacylated tRNA).

In some bacteria, uncharged tRNAs have been shown to bind to the ribosome. In fact, that binding is responsible for the "stringent response" a mechanism that signals that the cell is running low on amino acids1,2.
From Raina & Ibba (2014):

RelA is a ribosome-associated (p)ppGpp synthase which senses the presence of uncharged tRNAs that accumulate at the ribosome A site as a result of amino acid limitation. The presence of the uncharged tRNA acts as an effector molecule, stalling protein synthesis and activating RelA which then synthesizes pppGpp and ppGpp by phosphorylation of GTP or GDP using ATP as the phosphate donor (Haseltine and Block, 1973; Sy and Lipmann, 1973).

A similar mechanism has been proposed to operate in eukaryotes1,3,4.
Again, from the same section of Raina & Ibba (2014):

It has been proposed that discrimination between the charged and uncharged tRNA by Gcn2p occurs via an analogous mechanism of RelA protein activation as observed in E. coli by the presence of uncharged tRNA at the decoding (A) site on translating ribosomes. The activation of Gcn2p by uncharged tRNA requires its association with the ribosome via its C-terminal region and also, interactions between the N terminus of Gcn2p and the Gcn1p–Gcn20p protein complex which is also associated with the ribosome.

And, from the last paragraph of the discussion in Dong et. al. (2000):

We have argued previously that uncharged tRNAs bound to the decoding (A) site of the ribosome and base-paired with their cognate codons in mRNA activate GCN2 (15).

References:

Raina, M., & Ibba, M. (2014). tRNAs as regulators of biological processes. Frontiers in genetics, 5, 171.

Haseltine, W. A., & Block, R. (1973). Synthesis of guanosine tetra-and pentaphosphate requires the presence of a codon-specific, uncharged transfer ribonucleic acid in the acceptor site of ribosomes. Proceedings of the National Academy of Sciences, 70(5), 1564-1568.

Dong, J., Qiu, H., Garcia-Barrio, M., Anderson, J., & Hinnebusch, A. G. (2000). Uncharged tRNA activates GCN2 by displacing the protein kinase moiety from a bipartite tRNA-binding domain. Molecular cell, 6(2), 269-279.

Ramirez, M. A. N. U. E. L., Wek, R. C., & Hinnebusch, A. G. (1991). Ribosome association of GCN2 protein kinase, a translational activator of the GCN4 gene of Saccharomyces cerevisiae. Molecular and cellular biology, 11(6), 3027-3036.

So what is the answer to the question — why doesn't this binding disrupt protein synthesis? And where in reference 3 (or 1) is there anything about the activation of GCN2 being due to binding of deacylated tRNA to ribosomes?

@David — You're right, my answer kind of went off on a tangent — your answer is (as usual) excellent. If you're interested I added a bit more detail clarifying what I was basing my (non-)answer on. I've also clarified that the eukaryotic relevance is speculative. ——— I'll probably delete this later, but I want to think about how the "stringent response" and EF-Tu based discrimination work together ...

— A long ago I worked on tRNA binding to ribosomes, so I am familiar with some of the literature. There is a relatively recent paper on the stringent response that is relevant: Nature 534, 277–280 (2016). One recurring theme is the recognition of different structures and conformations of tRNA by different proteins, and I may add a section to my answer to point this out. In the case of stringent control one has RelA discriminating between acylated and deacylated tRNA bound to the ribosome.


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Does an antibiotic that inhibits protein synthesis delay bone generation?
After removal of a femoral intramedullary nail the bone needs to generate in the intramedullary canal and bone generation to take place at the screw sites. Would an antibiotic which inhibits protein synthesis, such as Clindamycin delay bone generation?

No, this cannot happen, because the ribosomes of prokaryotes (bacteria) and eukaryotes (mammals) are build differently. Ribosomes of prokaryotes consist of two subunits, the so-called 30S and 50S units, while the ribosomes of eukaryotes consist of one 40S and one 60S subunit.
Clindamycin works by inhibiting the 50S subunit of the bacterial ribosome and since inhibition is highly specific and the subunit is not present in mammals there is no inhibition of protein biosynthesis.

Mitochondrial ribosomes are similar to prokaryotic ribosomes and some antibiotics can indeed affect them. However, mitochondria are also "protected" because they are inside the cell. The linked paper also shows that mitochondrial ribosome is sensitive to some antibiotics but resistant to to others. (also this)


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What is this mushroom attached to the terrarium with transparent cap and black strips?
I have a terrarium with something black attached to the glass with a whitish green stem. Can someone please help identifying this?
The terrarium is outside in the backyard and receives only direct early morning sunlight -- it's currently summer. Location: Melbourne, Australia.
Terrarium internal side:

Terrarium external side:

Update:
Another exterior picture from the terrarium shows better details from the transparent cap and the black strips:

you are giving your succulents way to much water and you are giving us too little information.

@trondhansen, what information can I provide more than the photo?

what research have you done to try to find an answer to what this might be,where do you live,what temperature is your terrarium,what type of light do you use,for how long has this been growing in your terrarium.

@trondhansen, I have done enough research to accept being ridiculed at this website. The terrarium is outside at the backyard and receives only direct early morning sunlight. Here in Melbourne is summer.

this is very usefull information for the one(s)answering your question,please edit this into your question.and you should know that every single site here on SE want all of us to do some research before we ask a question.

I think (I'm definitely no expert) that this is one of the genus Parasola (Psathyrellaceae), they used to be part of a group that are known as inkcap mushrooms, though I see that this has changed. What gives it away is the thin cap and being able to see  the ribs in the cap looking like the ribs in a parasol or umbrella. It could also be one of the closely related Coprinellus or Coprinopsis, but these seem to have a thicker, less umbrella-like cap.
There are a number of species, and without access to spores and a spore print it would be impossible to tell which species it is.

I am searching more and found a term "Concave Translucent Mushrooms". Still not sure.

@ar2015 When I use those terms (on a different computer and network to when I wrote my answer) all I get is Coprinus (inkcap group - which used to include the Parasola) and Parasola genera coming up.


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what are the nodes and edges in gene regulatory networks
I am trying to find out how one can using gene expression data can infer gene regulatory network applying graph theory concepts. But I could not find a proper reference that
1)explain how one can get the adjacency matrix for a gene regulatory network?
2)if genes are nodes and edges are interactions between genes, where the gene expression values placed? and how one decide if there is an interaction between two genes?
3)what are the parameters there?
I found some literatures that describes DAGs but still the upper question are remained for me.
Any reference or help regarding these questions is highly appreciated if even it is going to be a keyword.

I think that this question is off-topic here as it does not relate to a specific problem in biology. You would do better transferring it to SE Bioinformatics to see if they will accept it.

You're asking for a short answer to an entire field of research. You won't get a comprehensive one that fits in a SE answer.
Generally, yes the genes/their products (which are often treated as the same thing in these abstracted models) are the nodes; the causal relationships between them are coded in (typically weighted, directed, bidirectional) edges. In graph theory more generally, you hold the "activity" of the nodes in a separate vector; sometimes that is taken as an input vector to your graph to produce a corresponding output vector at some time dT in the future (other times you aren't really interested in the activity at all and are working with just the graph itself).
Edge weights are determined by experiment, often many many interrelated experiments. If you experimentally increase or decrease expression of gene X, you can monitor gene Y for change to infer a "connection". However, it's more complex than that because an effect of X on Y could be mediated through a number of other nodes. Biology is rarely simple, and if you expect it to be you're going to make mistakes rather quickly.
If you have enough information about the graph and timeseries data about expression, you may be able to infer a model graph to describe the observed data. Some people have already compiled putative networks (particularly in model organisms) and you can find them by searching for their published papers describing them, or finding others who cite them.
If you want to do these things practically, I highly discourage working merely from textbook examples or broad summary reviews; to do something useful in science, you need to read primary literature. Your answers to these questions in your particular area of study will come by reading the methods sections of papers that publish on these networks.

Some references to start with:
Emmert-Streib, F., Glazko, G., & De Matos Simoes, R. (2012). Statistical inference and reverse engineering of gene regulatory networks from observational expression data. Frontiers in genetics, 3, 8.
Lee, T. I., Rinaldi, N. J., Robert, F., Odom, D. T., Bar-Joseph, Z., Gerber, G. K., ... & Young, R. A. (2002). Transcriptional regulatory networks in Saccharomyces cerevisiae. science, 298(5594), 799-804.
Pavlopoulos, G. A., Secrier, M., Moschopoulos, C. N., Soldatos, T. G., Kossida, S., Aerts, J., ... & Bagos, P. G. (2011). Using graph theory to analyze biological networks. BioData mining, 4(1), 1-27.
Vijesh, N., Chakrabarti, S. K., & Sreekumar, J. (2013). Modeling of gene regulatory networks: A review. Journal of Biomedical Science and Engineering, 6(02), 223.


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Why porifera is classified to have an incomplete digestive system although it has no organs?
Why are we using the term "incomplete" for digestive system only? We could have also said that for circulatory system that it is either incomplete or present as a simple system. What I mean is that, this doesn't relate to the definition of organ system. In other words, it will mean that the only organ system present in lower animals is digestive system.
Also for other organ systems, it is written that they are absent. What is the reason for this wording?

My advice would be to just learn what these labels are saying about the features the organisms have; the labels placed on them are not really important for "understanding the world", just for communicating with other people (like your biology instructors). Understand the system, then apply the label; don't use the labels to try to understand the system. The biology existed before the label.

An incomplete digestive system is one in which there is only one opening.  In our digestive system food is taken in by one opening (our mouth), the nutrients extracted and the remains are ejected by another opening (our anus).
In these lower organisms the food is taken in by one opening, the nutrients extracted and the remains are ejected through the same opening.
An absent digestive system is not having such a method of ingesting matter to be processed and ejected afterwards.  These organism absorb their required nutrients directly from the surrounding media.


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What is the optimal temperature for Omicron to thrive in?
I have always been under the impression that when sick, you should let a fever run its course. The accepted answer to the question Why do people take drugs to lower a fever? seems to provide some support for the theory and cites optimal temperature which says that most bacteria and viruses thrive best at 98.6 F. I was wondering how much variance there is to this optimal temperature and if some bacteria and virus infections can thrive at hotter temperatures? If so, I would think that studying the optimal thriving temperatures for various bacteria and viruses (including omicron) would be important to decide whether to treat, or let a fever run. With a quick google search I could not find any studies on the optimal temperature for omicron.

I think it's fine for this question to stand as a Biology question, but I think it asks the wrong thing if it's intending to use this as support for "should a fever be treated or not"; see https://meta.stackexchange.com/questions/66377/what-is-the-xy-problem and https://biology.stackexchange.com/a/57974/27148

Omikron is really new, don't expect comprehensive data on it. Even with SARS-CoV2, there is still much to learn and I am not aware of any "optimal temperature" studies (and I can't even imagine a design of such a study).


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How long is agar agar good for?
I'm a high school science teacher, and am starting at a new school. I'd like to do some labs with bacteria and so I'll need to make plates with agar agar. I found 3 huge jugs of powdered agar in the storage cabinet, but I'm wondering if they're still good. The only one with a date said "1995" and I think that may be a bit much, but I could be wrong. Does anyone know if I should buy new agar or use the old stuff?

There's a similar post on our sister-site for cooking: seasoned-advice.

I have used 5 year expired agar, no dramas. I expect your old agar will be fine.

I don't see why it shouldn't be ok as long as it has been stored dry. If it is still powdery and white it should be ok and personally, I would have any problems to use it in the lab.

I'm assuming these labs are going to be mostly qualitative in design? If you're able to use it to make an agar plate and grow bacteria on it, seems like it's still "good". If it won't form a gel then something's wrong. If it forms the gel and doesn't grow the bacteria... save that stuff and maybe consult with someone to find what new antibiotic you've discovered :)

Use science to figure it out!
Order some new jugs.  Tell the class about the old jugs you found.  Make 2 batches of plates: old and new.  Make dilutions of bacteria.  Does the old agar support as many colonies as the new?
Even better would be to have some other kinds of agar plates.  You can have old and new.  You can have agar with extra salt.  You can have agar with extra sugar.  You can have agar with a little iodine.  You can have agar made with old coffee.  You can have agar made with water from the fish aquarium.  Each student group will have a group of plates one of each type.
Then you can pool the plates of each type.  Graph number of colonies.  Use statistics.
If you determine the old agar is good then you can use it again next year and after that.  If you determine it is bad then it gets trashed.  Either way you are using the scientific method.  Yay science!

Test it first, but it should work fine
Agar Agar powder has a pretty long shelf life, even decades expired agar still works fine.
But if the container was opened it might have picked up some moisture from the air which will weaken how well it gels. It will be pretty obvious if you mix a test batch, just mix a bit in a paper cup and see if it gels. As long as it gels it will still work fine for any lab in a school setting. Any change will be uniform so if you used it alongside fresh agar there might be a noticeable difference in say PCR, but as long as you only compare it to itself even slightly bad agar will work fine.


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How can SNPs be well-defined if chromosome lengths differ?
I don't have a background in biology, but is trying to learn more about genetics; I have watched many videos about SNPs and still feel confused about the concept of a single nucleotide polymorphism. According to wikipedia, a SNP "is a germline substitution of a single nucleotide at a specific position in the genome". What I am most puzzled by is the concept of a 'specific position in the genome', if chromosome lengths differ, how can we talk of a 'specific position'?
For example, if we have two arrays of items that are of the same length:[A,C,C,C,A,T,G] and [A,C,G,C,A,T,G] then it is straightforward to that the entries at the third position of each array are different. But with chromosomes, their lengths differ, so it might look like: [A,C,C,C,A,T,G] and [A,C,C,C,A,A,T,G], now how can we say what corresponds to what? Do we say there is a SNP at the fifth/sixth base pair?
What about something more different like [A,C,C,C,A,A,T,G] and [C,C,A,A]? How do we know what corresponds to what? What are the SNPs?
What if two individuals each has a chromosome 1 that differ by many base pairs, how do we know what corresponds to what?

The reason we can compare SNPs between different people is because sequencing reads are aligned to a reference genome with a fixed set of coordinates for each chromosome (local alignment), rather than aligning the entire chromosomes of two different people (global alignment). For instance, 10:3424234 refers to the 3424234th base-pair on chromosome 10 on the reference genome.
Say we wish to compare this particular SNP between two different people; we can sequence both individuals and reads covering the 3424234th base-pair on chromosome 10 will be produced and aligned to the same part of the reference genome; we are certain that the reads originate from the same part of the genome in both individuals. Then, it's trivial to look at this position and see whether it differs between the two individuals. It therefore doesn't matter if one individual has chromosome 10 twice the length of the other (although this would never be the case in reality), as long as the read originates from the same part of the genome in both individuals, they can be compared.

I have accepted this answer although I don't fully understand it. Could you please tell me where I can find out more how "reads covering the 3424234th base-pair on chromosome 10 will be produced and aligned to the same part of the reference genome"? Any textbooks recommendations?

In the image, say that we are interested in base pair 114. Read2 is aligned to that position and tells us the genotype at this position. If we do this for two people, then we can compare the sequences on the reads to see if they are the same or different. Does that help?

Please allow me to ask 2 different questions: How do we know that Read1 is correctly aligned? There are 2 discrepancies. How do we know that Read3 is correctly aligned? Is this all done by statistical/computational methods?

Yes, the reads are about 25 base-pairs long, and there are sophisticated algorithms to make sure they are aligned correctly, e.g. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2705234/

Read3 has two extra C's in the middle, does this imply this person inherited a mutation (or two mutations), specifically, 2 C insertions? It looks like this procedure works because human genomes are largely the same.

Yes - exactly right on both counts.

It may be easier to understand this with an analogy. Think of the genome as a book, and the chromosome as a chapter in the book.
If someone were copying the book by hand (as was common before the invention of the printing press), they'll make some mistakes. The vast majority will be copied correctly, but here and there they'll misspell something, and might even occasionally leave out of duplicate a word.
These mistakes are analogous to mutations during genome replication. As long as they're rare, you can easily recognize all the common parts of the original and copy, and the places with the errors stand out as SNPs.
E.g. if you go from

The quick brown fox jumped over the lazy dog

to

The quicck brown fox jumped over a lazy dog

you can see that an extra "c" was added in the second word, and the second "the" was replaced with "a". But for the most part the two sentences are the same. And just as we're able to make sense of the sentence with these errors, the genome has enough redundancy (as well as "junk" that has little effect) that the genome still works with a few errors and might even be beneficial (but some can cause diseases, or be catastrophic enough to prevent the organism from developing at all).
SNPs aren't the only kinds of mutations, they're just some of the easiest to analyze because they stand out so well. For instance, humans have 46 chromosomes, while great apes have 48. Scientists have determined that human chromosome 2 resulted from the fusion of two chromosomes from the common ancestor of humans and apes. The analogy would be an editor deciding to combine chapters of the book when releasing a new edition (although in evolution there's no editor making conscious decisions, it just happened accidentally, as if the chapter heading was left out when copying). It took some genetic sleuthing to determine whether this was originally two chromosomes that merged in humans, or one chromosome that split in apes; you can read about it here.

Agreed this is a good analogy. Some other aspects that the analogy helps with is showing why you need context; if you just had the word "quick" it would probably be hard to ID the typo as just a "SNP", just like in OP's 4-7 base examples (maybe the original was "duck" and both the q and i are inserted). But, once you have the whole sentence, it's clear where exactly the difference lies. It also doesn't matter if you only started copying from Chapter 3, or have extra introductory pages; it's still clear where the exact change is from the local context, regardless of the total length.

Good analogy, thank you.

But as more and more mutations occur (like in 500 million years), the genomes will become less and less comparable, right?

@NoppaweeApichonpongpan Yes, although there are still meaningful similarities. There are some genes that control some basic biological processes that are common to very disparate species. It's kind of like the way most fairy tales begin with "Once upon a time".

Or the way each chapter of a book begins with "Chapter ##".

so it might look like: [A,C,C,C,A,T,G] and [A,C,C,C,A,A,T,G], now how
can we say what corresponds to what? Do we say there is a SNP at the
fifth/sixth base pair?

We say the second one has an insertion at 5.
Have you read the vcf format?  It describes how all kinds of variants are documented.
https://samtools.github.io/hts-specs/VCFv4.2.pdf

What about something more different like [A,C,C,C,A,A,T,G] and
[C,C,A,A]? How do we know what corresponds to what? What are the SNPs?

Absent any context, why would you want to compare two completely different sequences like that?
If you really wanted to compare them, surely single nucleotide polymorphisms isn't the right way to think about the differences.

I don't yet have a clear picture of what an organism's genome is 'supposed' to look like. Thank you for your answer.


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Seed head Identification. July in Toronto, Ontario, Canada
What flower has this type of seed head? (this was shot on July 17th, 2020)
This was in a residential garden in Toronto. Going from memory, i believe the size was approx. 2 1/2" to 3" in diamater.

Looks like an allium to me, but I doubt anyone is going to be able to identify it more specifically from only this part of the plant - there are hundreds of alliums, both cultivated and wild. Do you have other photos of the rest?

Thank-you Bryan, that's really helpful. This is the only photo i have.

Alright; I'll post it as an answer. Possibly someone else will be able to narrow it down a bit more for you.

Looks to me like an allium, but you'll find many of these look quite similar at this stage, I doubt you can get much further than that. See for example a Google image search for allium+flower+seeds (added 'seeds' to the search to get later stages like the one you've got).
Alliums include onions and garlic (both cultivated versions you'd commonly find in a grocery store and wild varieties; for onions this includes green onions and scallions as well as bulb onions), numerous other plants grown primarily for decorative purposes commonly called "allium", and lots of other wild plants.

I agree with @BryanKrause that this is an Allium seed head. The seed pod and seed on the end give it away
For another partial match from a commonly cultivated plant in domestic gardens, Agapanthus seed heads look similar, though the seed pod and seeds are elongated and winged. When these dehisce at the end of seeding you can be left with a head that superficially looks like OP's photo, but without the small black seeds. Agapanthus tend to be much larger, up to about 6 inches (15 cm) across, though there are many cultivars, some of which have smaller inflorescences.
Head with seed pods (from gardeningknowhow.com, photo by eldadcarin):

Without pods (from seenobjects.org, photo by Martin Kenny):


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Hypothalamus and posterior pituitary gland in ADH production
Concerning the role of hypothalamus and the posterior pituitary gland in osmoregulation and especially when water potential of the blood is low,
When the osmoreceptors in the hypothalamus detect the low water potential of the blood, they lose water by osmosis and shrink, my question is:
NUMBER 1 》does this shrinkage stimulates the syntheisis of the ADH in the neurosecretory cells in hypothalamus first and then ADH is sent along the axons where they terminate in the posterior pituitary gland secondly and released into the blood stream thirdly (just non stop) , OR
NUMBER 2 》 before the shrinkage happens, already ADH was synthesized in the neurosecretory cells and was sent along the axons where they terminate in the posterior pituitary and ADH was stored there in the axon terminals and whenever the neuroscretory cells shrink, impulses are sent along the axons stimulating the release of ADH from the posterior pituitary gland into the blood stream?
Because I have read both explanations from different resources and I am confused which is the correct physiology mechanism that releases ADH from posterior pituitary gland.

Your second hypothesis is correct.
Hypothalamus secretes vasopressin (ADH), transports it along the axons and stores it in the posterior pituitary, thereby when hypothalamus senses the stimuli(osmotic potential or H+ levels or some other indicators) it generates nerve impulse and sends along the posterior pituitary to release that stored ADH.
Comparison of Both Cases
To get insight into the answer lets consider both hypothesis one by one and compare them on the basis of speed of action, in the first case as you mentioned if hypothalamus starts to transport ADH on stimuli this will lead to slow effect (and a comparatively long time / lag to show effect) also once if hypothalamus feels it needs to secrete more ADH since it wasn't enough to meet the requirements it will take even longer time which could be a disadvantage to the mechanism overall. Whereas in second hypothesis these limitations are take care of - hypothalamus transports it to Posterior pituitary from where it can deploy the effect quicker than earlier situation, also it can store some extra ADH in it's own once it has bulked the posterior pituitary's potential which is useful as a secondary backup.
Supporting Citation:https://pubmed.ncbi.nlm.nih.gov/2046874/

Please note that speculation is not welcome on this site, particularly on subjects for which research is available — consequently your answer is much more likely to receive a favorable response if you edit it to include supporting references (primary literature is best). Without that support, your answer is indistinguishable from opinion and thus is inappropriate for this site. ——— Please take the [tour] and then consult the [help] pages for additional advice on [Answer] effectively on this site and then [edit] your answer accordingly.

That speculation was from the mind which is well learnt to speculate upon that. Also it is not possible to provide for the citations in every kind of question which can be well defined with some dogma. Thanks for the concern.


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Weird spiky bug found in northern Spain
Does anyone know what this bug is? We're located in the north of Spain. Around 5mm long with spikes in its back.
First image is belly up, second one the bug is walking.

The second photo looks almost exactly like the first, except blurry and without a shadow. Is that a mistake? Can you take a clearer photo of the dorsum (back) of the bug? Closer would be great! Thanks.

Sorry, they're not mine and the bug is no more. Supposedly the second one is the dorsum of the bug. I'm another site someone suggested it found be a cordyceps infected bug (didn't know the bug's species though :(

OK, I'll assume that because there are no legs visible adjacent the surface where there should be, this is just another picture of the upside down bug. The chances that there would be such a symmetrical multiple cordyceps infection are remote, to say the least.

Oh, then back to square one  it's a shame there aren't better pictures.


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How is the extraordinary eyesight of some deep-sea creatures explained?
Mantis shrimp is a marine crustacean that is known to have a vision system that is much more complex than the eyes of many animals living on the ground. As explained here, it has 12 types of photoreceptor cells, can detect a wider frequency range than humans (into UV and infrared regions), and even polarization.
These animals live in dark underwater environments where sunlight cannot reach, so that a vision system would not seem to be useful.  Why is this present?
My own thoughts are that even though some types of these creatures live in deep ocean waters, they mostly live in shallow waters where sunlight can reach:

"They occur in coastal waters but are sometimes found as deep as 1,300 meters"

Therefore their vision system is not useless. But what about those species (including but not limited to some types of mantis shrimps) that live in deep dark waters? Is their vision system considered a vestigial feature?

It's an interesting question, but could you [edit] to make the title consistent with the question in the body?

@ARogueAnt. I edited the title to better match the question. Feel free to edit it!

I have edited your question to reduce it to its essentials. This is “what purpose does this serve?”  If you can answer that you know why it evolved. If you can’t, you are in trouble.

Its worth noting that mantis shrimp eyesight is much worse than ours in some way, they are far less sensitive to small differences in color while detecting a much wider range of colors. it is also worth asking if deep water mantis shrimp stay in deep water all the time.

You will find quite a lengthy and nice discussion and review of current hypotheses (all with some support) in Porter et al. 2020,PNAS.
To add even more intrigue, parts of the vision system (e.g.: number of photoreceptor genes) seems more complex than needed for the actual performance of the visual system (e.g.: whether certain visual stimuli are detected).
Suspected:

Parts of "visual system" do not primarily serve "vision" but something else (e.g.: touch as in some insects)
There is an advantage for heterozygous animals
There might be some sort of vision-based advantage to be discovered (e.g.: rather than helping to better discriminate seen things in real time, the complexity might serve to integrate visual cues over time)


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To which thing does "two-thirds" refer here?
Life and Letters of Charles Darwin, volume 2 includes the following letter from Darwin to Hooker, about a examining a plant that absorbs ammonia:-

Reflect how little ammonia a plant can get growing on poor soil—yet it is nourished. The really surprising part seems to me that the effect should be visible, and not under very high power; for after trying a high power, I thought it would be safer not to consider any effect which was not plainly visible under a two-thirds object glass and middle eye-piece.

To which thing does "two-thirds" refer here, i.e the size or magnification or something else?

For future reference, it looks to me like this type of question would be more appropriate on [HSM.SE].

The use here of "object glass" is equivalent to our current usage of "objective lens" on a microscope. In this case Darwin used a 2/3 lens and the "middle eye-piece" from a particular microscope. I think (but can't find confirmation) that the 2/3 refers to the width of the field of view in inches.
The letter you quote is to Edward Cresy Jr. on the 2nd November 1860. I can't find much on Darwin's microscopes, other than in the Whipple Museum, where they have 2 microscopes of Darwin's. One of these purchased in 1847 is a microscope with exchangeable lenses. This microscope was a Smith and Beck compound microscope. You can find examples of these for sale in antiques collections and a similar one is probably this one, which comes with a number of objectives and a 3 magnifications of eye-pieces (~1/2 way down page). The magnification of a 2/3 objective looks to me to be about 10-20x (confirmed in this review as ~15x (about 2/3 way down page)).
As for the middle eye-piece, based on the same  Microscopy UK page linked above, it is probably a #2 eye-piece of about 8.5x, so a total magnification of about 125x (127.5x, if the numbers are accurate).


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Issues obtaining Fo in DIY Imaging Fluorometer
I have successfully built an imaging fluorometer at home (see project details here) and so far is working amazingly well.
But I am having serious issues in obtaining Fo to calculate Fv/Fm (Fm-Fo)/Fm. The reason is less technical than theoretical, as after scouting dozens of academic papers all I can find are contradicting accounts as to how to set up the fluorometer to effectively obtain an Fo value.
Without Fo, the Fv/Fm values of all plants I have analysed range around ± 0.90 which is way above the usual 0.83 the literature cites as an unstressed plant. If the number were below 0.83, I would believe the plants are stressed, but a higher number clearly indicates my setup is not correct.
Most literature states that the measuring beam must be of the order of < 1 μmoles m−2 s−1 to avoid eliciting any photosynthetic rate. The reason for the literature to be so scant about this protocol is that 99% of fluorescence research is done with commercial fluorometers and researchers don't need to adjust parameters as they are adjusted by the software. There seems to be a bit of esoteric mistery around these fluorometer parameters, probably due to some commercial sensitiveness.
Under my setup, using a measuring light of < 1 μmoles m−2 s−1 elicits a signal of 0.3 on the camera, while the saturating pulse can be over 100 (0-255 scale or 8 bits) and above depending on intensity. Calculating Fv/Fm using these numbers gives a number above 0.98 which is clearly wrong. Increasing the camera exposure does not help.
Another issue is that we are talking about fluorescence imaging and not the traditional photodiode based fluorometer which is closer to the source and way more sensitive. A camera is far away from the subject and increasing too much exposure leads to noise, etc.
Then again, I do not think this is a technical issue, as I've got the feeling the measuring light of the commercial imaging fluorometers is much stronger than otherwise stated.
Has anybody dealt with this specific issue?

This paper1 gives a definition of all the various parameters. F0 (that's Fzero, not F letter "o" I think) is defined as the dark state fluorescence. There is a second parameter F0' that is the light state fluorescence.
F0 would be your ground-state with minimal induced fluorescence, it seems that this might require "overnight in the dark" for true measurement values. Measure the fluorescence with the measurement (far-red I think) light turned on. You should then apply a saturating pulse of actinic light to measure FM. You should also determine the background from your rig using reads taken with no sample in place and with and without measurement light turned on. This paper2 provides a description:

The calculation of fluorescence parameters is probably best explained by reference to a typical experimental trace (Fig. 1). Under field conditions, an experiment of this type may not always be appropriate, however, most of the parameters can still be calculated. The measurement is initiated by switching on the measuring light, giving a measure of the Fo (minimal) level of fluorescence. A saturating flash of light is then applied, allowing the measurement of Fm in the dark‐adapted state (
Fom
). Following on from this, an actinic light is applied and, at appropriate intervals, further saturating flashes are applied. From each of these, a value for Fm'
, the fluorescence maximum in the light, can be measured. The steady‐state value of fluorescence immediately prior to the flash is termed Ft. After a flash, removal of actinic light (preferably whilst simultaneously giving a far‐red light) allows measurement of Fm'
.

1: Lichtenthaler et al. Photosynthetica volume 43, pages 379–393 (2005)
2: Maxwell and Johnson. Journal of Experimental Botany, Volume 51, Issue 345, April 2000, Pages 659–668

I am well aware of those papers and they don't answer the fundamental question, nor does your answer. In the meantime, I have found the answer to my question but I need to test it before posting my answer here.

@MaykeSantos - I read your question to ask how to set F0, due to this bit "But I am having serious issues in obtaining Fo to calculate Fv/Fm (Fm-Fo)/Fm." I don't see any other questions in your post (other than last sentence). If you could solve F0, then your problem would disappear. Can you calculate F0' as a proxy?  the literature suggests that this is a reasonable substitute. I doubt the measurement light on commercial instruments is strong as calibration is a common part of set-up when purchasing these sorts of instruments.

yes, the question is how to calculate F0 in a Chl imaging fluorometer system, that is different to most common photodiode-fibre fluorometers. There is a long paper about how to calculate F0' (not F0) as a proxy, but that's different and it can be achieved with far red light. Now, to answer the question itself, my problem is light modulation. I need to apply a very strong light pulse but at a much higher frequency (below 300us) in order to avoid eliciting photosynthesis. The answer is in Oxborough 2004

@MaykeSantos - you should edit that into your question, as it isn't clearly stated what you are actually looking for. 300 us is very short, you might be able to do it with a shutter mechanism, but you would need a very fast processor controller. You might be better asking for migration to the electronics SE

I will leave it as it is. Thanks.


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Where does nitrogen go or come from as its partial pressure in different parts of the lungs decreases or increases?
Here's a table from Guyton and Hall Textbook of Medical Physiology showing the partial pressures of gases in different types of air:

We can see that as the partial pressures of some gases decrease or increase, the partial pressures of the other gases increase or decrease, respectively. Let's focus on nitrogen. The PN2 in humidified air is lower than in atmospheric air because the PH2O in humidified air is higher (I'm not sure if the word "because" is correct here, but I guess it is). The PN2 in alveolar air is higher than in humidified air because the PO2 in alveolar air is lower.
Since partial pressure is proportional to concentration, I believe a gas should go somewhere for its partial pressure to decrease or come from somewhere for its partial pressure to increase.
Where do nitrogen molecules go or come from in the two cases I described? As the PN2 in humidified air decreases, where do the nitrogen molecules that cause this decrease go? As the PN2 in alveolar air increases, where do the additional nitrogen molecules that cause this increase come from?
I'm asking only about nitrogen because I think the reasons should be similar for the other gases (i.e., the PO2 in humidified air and atmospheric air) or are clear (i.e., diffusion into and from the blood for O2 and CO2).

You can't just look at nitrogen and how its concentration changes. Their concentrations all rely on each other. Nitrogen is inert; in your examples it doesn't go anywhere. The partial pressure of N decreases  in humidified air because more water is in the air, making the percent nitrogen fall. The N in inspired and expired air is basically unchanged. We add a bit of moisture to exhaled air.

You can get an answer in the accompanying text in Guyton & Hall that describes your table. It says: "Because the total pressure in the alveoli cannot rise to more than the atmospheric pressure (760 mm Hg at sea level), this water vapor simply dilutes all the other gases in the inspired air. Table 40-1 also shows that humidification of the air dilutes the oxygen partial pressure at sea level from an average of 159 mm Hg in atmospheric air to 149 mm Hg in the humidified air, and it dilutes the nitrogen partial pressure from 597 to 563 mm Hg."

@anongoodnurse If nitrogen didn't go anywhere, wouldn't that increase the total pressure? For example, imagine a container that contains only gaseous nitrogen. This nitrogen creates some pressure. If we added some water molecules, these molecules would too create some pressure. If we didn't remove some of the nitrogen, I think the total pressure would increase, since it would be a sum of the pressures created by the nitrogen and water.

@Domen I saw this explanation and a similar one in Costanzo's textbook, but I didn't understand both of them, unfortunately. In the case of liquids, when you dilute something with water, you increase the volume of the solution. This makes me think the volume of the gas in the lungs should also increase. Sorry, I'm not very good at pure physics.

@Domen I think you should write that as an answer.

@athlonusm Your two comments are related: the way it wouldn't affect the total pressure would be by affecting volume instead. In this case you really shouldn't worry about pressure at all though, just assume it's always the same; it's forced to be the same by the measurement approach, anyways.

@athlonusm - This isn't as much about physics as about straight math, or if you must, algebra. For example, if you have a 900 cc of nitrogen in a big floppy balloon (so, 100%) and you add 100 cc of water. The amount of nitrogen doesn't change, i.e. the N didn't go anywhere, but its concentration dropped to 90%, and H20 goes from 0% to 10%. (<-- that's a tree). You're looking at a tree, when you should be looking at the forest (all the components and how they relate to each other causing percentages of each to change when one component changes.)


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What does the 'y' mean in CHL1−/− / L1−/y double mutant mice?
I am reading a journal paper and I have come across the following statement:

To investigate this, $CHL1^{−/−}$ / $L1^{−/y}$ double mutant mice were generated
and analyzed for thalamocortical axon topography.

I know that -/- means knockout, +/- means heterozygous and +/+ means wild-type. But what does -/y mean? I have never seen this notation before.
Any insights are appreciated.

According to the introduction, human L1 is located at Xq28, in the q arm of the X chromosome. It is also on the mouse X chromosome. The $CHL1^{-/-}$ nomenclature you've already figured out: a homozygous knockout of the CHL1 gene. I haven't thoroughly read the whole paper, but there must be some undesirable phenotype when L1 is homozygously knocked out in female (X/X) mice, so the researchers in this paper are using male (X/Y) mice, with L1 knocked out on the single X chromosome. There is no corresponding allele on the Y chromosome, so the nomenclature is $L1^{-/y}$.
This is borne out in the Materials and Methods:

The L1 gene is located on the X chromosome, and its deletion results in poor breeding capability of males and thus female mice homozygous null for the CHL1 gene and heterozygous for L1 ($CHL1^{-/-}$/$L1^{+/-}$; C57BL/6/Sv129, ∼9:1) were crossed with $CHL1^{-/-}$/$L1^{+/y}$ males (C57BL/6) to produce $CHL1^{-/-}$/$L1^{-/y}$ double mutants.

From their methods as you've quoted it seems that the issue is that the male L1 knockouts don't breed well; that makes it difficult (perhaps impossible?) to produce a female with both copies knocked out, because one of those has to come from the male-that-doesn't-breed-well. There may not be an known undesirable phenotype for the female double knockouts, just that there's no way to produce them by breeding, and it's much easier to just genotype the male offspring.

I read that section from the M&Ms multiple times, and I still couldn't figure out exactly what the issue was. You finally broke through my brain fog, thanks :)

Took me a second too... my first thought was they were just using male mice for other convenience reasons; it's been fairly common in neuroscience to use only male rodents to reduce variance in measurements from sex differences and estrous cycle, though there's a bit of a push against that trend these days.


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How long could Jack Sparrow and Will Turner survive underwater with a row boat over their heads (Pirates of the Caribbean I)?
How long could Jack Sparrow and Will Turner survive underwater with a row boat over their heads (Pirates of the Caribbean I)?
This scene would be physically infeasible due to buoyancy issues (http://kwc.org/mythbusters/2007/11/episode_92_pirates_2_row_boat.html). One may try to solve this by loading the boat itself with some heavy weights.

Here, the question is about the physiological feasibility - how long they can last with the air in that row boat. The question occurred to me because I have seen people in forumns claiming 2 people could not have made that journey with the air present in that space. But I don't think this is an issue.

+1 2 people downvoted your post without any reason and your question is fine and I think it is very interesting.

This is my attempted solution that I would like to share and verify.

Volume of air.
Assume dimensions of the boat (modeled a cuboid) = 4m x 1m x 0.5m
Thus Volume of air, V = 2 m3.

Acceptable limits of air composition
Carbon dioxide (CO2) poisoning will happen far before there is any risk of running out of oxygen (O2).
Referring to https://en.wikipedia.org/wiki/Carbon_dioxide#Toxicity & https://en.wikipedia.org/wiki/Hypercapnia#Tolerance, I am taking up to 5% concentration of CO2 as acceptable for the duration of the underwater walk.

Actual air composition calculation
3.1. Initial air composition (https://en.wikipedia.org/wiki/Atmospheric_chemistry#Atmospheric_composition):

O2 (initial) = 21%
CO2 (initial) = 0.04% (or 400 ppm)

Thus, for 2 m3 of air (= 2000 L),

O2 (initial) = 420 L
CO2 (initial) = 0.8 L

3.2. Final air composition (Based on acceptable limits above, final concentration of CO2 = 5%). Thus:

O2 (final) = 16%
CO2 (final) = 5%

Thus, for 2 m3 of air (= 2000 L),

O2 (initial) = 320 L
CO2 (initial) = 100 L

Thus, we have time until the 2 people (Jack and Will) have exhaled 100 L of CO2.
3.3. Time taken to exhale 100 L of CO2 (by 1 person).
Composition of exhaled air (https://en.wikipedia.org/wiki/Breathing#Composition):

O2 (exhaled breath) = 17%
CO2 (exhaled breath) = 4%

Volume of exhaled air per breath (https://en.wikipedia.org/wiki/Breathing#Gas_exchange) = 350 mL.
Thus:

O2 (exhaled breath) = 17% of 350 mL = 59.5%
CO2 (exhaled breath) = 4% of 350 mL = 14 mL

Thus, Volume of exhaled CO2 per breath = 14 mL
Assuming 15 breaths per minute,
=> Volume of exhaled air per minute = 15 * 14 mL = 210 mL
=> CO2 exhalation rate = 210 mL / minute
Thus time taken to exhale 100 L of CO2 (by 1 person) = (100 L) / (210 mL / minute)
= (100 * 1000 mL) / (210 mL / minute) = 476 minutes.
=> time taken to exhale 100 L of CO2 by 2 people = 476 / 2 minutes = 238 minutes ~ 4 hours.

Thus, Jack and Will will be able to last 4 hours breathing from that air under the row boat before the air concentration of CO2 reaches 5% (and O2 reaches 16%).

You've not taken into account that the CO2 dissolves in the water, the surface area of the water, the rate of dissolution. Also... maybe additional O2 from the water? (I'd need to check that). That would need to be balanced-out against extra metabolic and muscular use of oxygen from the effort expended.. Just a thought.

@ARogueAnt. As long as the pO2 is different in the water and the air there will be a net diffusion between them. pO2 of the water will be roughly equivalent to atmospheric pO2 at the surface, where air and water are mixed.

Glad you took the time to answer your own question. It's a well-laid out answer. Good job! I agree with the previous comment about CO2 (though I'm less convinced the oxygen pressures will make a big enough difference to matter). Anyway, the reason for my comment is that I wanted to additionally point out that 15 breaths/min might be reasonable while at rest, but these two individuals would likely be breathing faster (i.e., producing more CO2 per minute) given their activity. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4818249/ suggests 40-60 breaths/min is more normal during exercise.


text:
Is it possible to identify secreted metabolites or secondary metabolites from a gene list?
So I have a gene list obtained from differential gene expression analysis of a metabolic disease model, and I was wondering if I can identify metabolites or secondary metabolites that may be produced by some of enzymes or reactions modulated by these genes by bulk.
For example, I know that Ptgds synthesises prostaglandin D2 and it is secreted in urine and plasma, and so on.
However, I don't want to do a whole research on each and every gene separately, rather like a database or a tool that can identify them by just inserting a gene list!

Obtaining the specific metabolites from genome-wide measurements of transcripts can be somewhat possible, but remains quite challenging, and might not be possible for many organisms because too little is known about them. If the purpose is to get a better idea of the changes in transcriptome without needing to infer specific metabolites (e.g.: if it is sufficient to know if some pathway is up or down regulated) you could do functional gene enrichment analysis instead. Latter can be adopted widely and there are nice tools such as the web-based https://david-d.ncifcrf.gov

The problem is that we want to identify some biomarkers that are steadily found in urine, plsama or other easily attainable fluids. I already did enrichment analysis and gene ontology along with many others.

I am currently going through each gene and looking into their pathways to find whether they produce metabolites. It's time consuming and not efficienct I'm afraid.


text:
Could hydrogen peroxide (H2O2) be used as a 'disinfectant' on hospital patients that are in contact with SARS-CoV-2?
Hydrogen peroxide kills harmful pathogens and various forms of bacteria by destroying their cell walls, through a process called oxidation. However, since SARS-CoV-2 is acellular; not composed of a defined cellular structure, would hydrogen peroxide still function as an antiseptic in this case?
I am aware that there are other forms of cleaning and disinfection products which has being proven to kill SARS-CoV-2, however, I am curious to understand the practical applications of hydrogen peroxide.

i did a google search(you did not)you can take a look at part 7 here to find an answer to your question https://en.wikipedia.org/wiki/Hydrogen_peroxide.it is expected that you do a minimum of research before asking a question,a more focused question will often get you better answers.

Check out the "List N" from the USA CDC and check out the chemicals used for disinfection that have been tested and found effective against SARS-CoV-2.
Hydrogen peroxide (H2O2) can be used as an antiseptic at low concentrations, but would generally not be used in a clinical setting for control of a virus. It would generally be used as a treatment for cleaning of minor wounds, but is no longer considered to be a suitable cleaning agent for this purpose because of irritation of the skin that happens as a result of application. It can not be used internally.
High concentrations (over about 5%) can produce severe burns on skin and other organs by inhalation or ingestion. Proper PPE used correctly is essential when using these products.
You can, however, as the list N suggests, use it very well as a cleaning agent. It is very bleaching, so it is not a good idea to use it on clothes or materials that are darkly coloured without testing them first.

High-concentration $H_2O_2$ can also produce sometimes severe burns on unprotected skin, as anyone who has performed immunohistochemistry has probably learned...

@MattDMo - yes, I think I should have stated that clearly, rather than by implication - will edit in.


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Why there is swelling when skin doesn't break visibly?
I had read that swelling occurs as result of blood vessels pouring watery fluid in the affected area to help in fighting the incoming pathogens, when we our skin is cut open. However sometimes like stubbing toe, or getting hit in head, our skin doesnt breaks, but still swelling occurs on the affected region. Whats the point of swelling in that case, when pathogens obviusly are not invading, as our skin is not cut open?

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text:
What's the best way to find a Tardigrade
So I have a school science project on the best sample to look under a microscope, so I want to bring in a Tardigrade. It's winter here in Ohio, although Tardigrades are freeze tolerant I don't think there's a high chance of finding one in a cup full of snow. I heard Tardigrades live on certain fruits such as strawberries and lettuce. I just want to be 100% sure I'll bring in a Tardigrade.

P.S: It's snowy so I can't make a tardigrade habitat.

google where to buy tardigrades,you can get them online

Have you searched for where Tardigrades usually live? (should be easy to find). Please do so (since we require question askers to demonstrate some research effort, and a simple Google or Wikipedia search would likely give you a couple of starting points). Update your post with any updates and where you found the info, and then let us know what you still can't figure out. Thanks

Alright, Thanks.

If the ground wasn't frozen I would have not posted this question.

I have heard they live in moss, so one time I got some moss and tried to find tardigrades in it. This did not work. I upvoted your question because I would love to find tardigrades too. It's not fun to "get them online."

Maybe greenhouses? or Houseplants? Indoor botanical gardens (which is basically a greenhouse in museum form)? You could ask the staff there if you have one.

Alright, thanks for the help everyone.


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Western blot trouble shooting - low voltage and yellow sponges during/after transfer
I have a western blot troubleshooting question that I haven't been able to find the answer to in manufacturer troubleshooting guides.
As a bit of background, I was transferring 2 western blots together in the same tank (we use a BioRad tank that fits 2 transfer modules side-by-side). Normally when transferring two blots in the same tank we run for 2hrs at 20v, as the tank takes about 30 minutes to get up to 20v. However in this case after 1hr the blot was still only running at 4v. I checked the tank and the power pack temperature, and both were not noticeably hot, so I don't think the pack was struggling with high resistance. After 2h the blot was still only at 4v, and had barely transferred at all.
Upon dis-assembling the transfer module I noticed that the sponged inside all had turned yellow, however only the sponge touching the positive electrode, and only the side of the sponge facing the positive electrode.
I'm not really sure what went wrong here to be honest. The only thing that I notably did differently was that I made the transfer buffer up the day before and left it in the ice machine overnight. The transfer buffer was glycine and tris with 20% methanol, in distilled water.
Any thoughts would be appreciated.

What concentration of glycine in the buffer? Sound very much like your transfer buffer wasn't correctly made or you had a non-conductive layer in the western that wouldn't normally be there. It is very unusual for Voltage to not reach the set point almost immediately.

Did you check the wiring in the transfer tank? Something could be corroded or otherwise damaged to the point of breaking, allowing only weak current to pass through. Also, make sure the leads are clean (use a brass brush to remove any corrosion) and of course check that your power supply is working properly...

Thank you for the comments. When I got back into the lab I double-checked the chemical shelf and it turns out I had put NaCl into the buffer instead of glycine! Currently soaking the sponges over the weekend to see if the yellow colour comes out...


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What is the shortest read length achievable using Oxford Nanopore sequencers?
Is there a lower limit, in terms of read length, one could expect from sequencing DNA on Oxford Nanopore devices?

It depends on the size of the DNA fragments you are sequencing. They use 400bp amplicon libraries for their SARS-COV-2 sequencing protocol, which is about the shortest library prep I've seen for Nanopore. If your DNA is highly fragmented, you can probably expect a lot of reads around the 500 bp range or smaller.

I've had fragments come out in the low 10's of  bases - using non-standard protocols in a non-standard way, where you can see read through of both adapters.

@bob1 low 10’s including the adaptors?

Do you know if it is mentioned in the manual or in the sequencers specs or else? I would love to have a reference.

@alec_djinn no, actual sequence. In that case the adapters were (IIRC) about 50-80 bp each, though I would need to re-visit the sequences to confirm. I guess that total sequence length of ~150 bp as minimal. Though that would be the ones that passed QC, I haven't looked at the ones that didn't.


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What protein(s) serve as the control antigen in Covid lateral flow tests?
Having recently been obliged to take a lateral flow test for Covid in order to fly from the UK, I idly wondered:

What is the control antigen for Covid lateral flow tests?

as (fortunately) that was the only band I saw.
Easy — just Google? You jest! “Other people asked”, “All you need to know” and even the scientific papers linked just talk about the target antigen.
This sort of thing really gets up my nose!
Could someone please tell me what it is up my nose?

You mean the red stripe at the upper control position of the test? See this question where it has been answered: https://biology.stackexchange.com/questions/101767/whats-happening-in-the-c-and-t-stripes-of-a-covid-test-kit

@Chris. I see. It is the test antibody itself. So this would seem to control only for addition of the test solution, and not whether I’ve swabbed my nose adequately. Is this correct? If so, it doesn’t seem correct.

@David, you are correct. Even if you don't swab the nose and put "clean" extraction buffer on the test card, you will get the "C" band. On my tests, even tap water gives the "C" band (although a bit lighter, but still clearly visible).

Absolutely correct. There is no control if you took the specimen correctly. Luckily most "corona skeptics" do not understand how this test works. As @Domen says, you get the control band with water only. Oh, btw. you can also fake a positive best by using a few drops of apple juice or coke.

@Domen Thanks. Well I’m waiting for a plane to take me to the ski slopes now. No point in going if I didn’t really test negative. First time in an airport for two years — never seen it so empty. Trust I can negotiate Swiss and French controls.


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Is the volume of blood pumped by the right and left ventricles the same?
Wikipedia says that the definition of stroke volume is:

stroke volume (SV) is the volume of blood pumped from the left ventricle per beat

Why the left ventricle specifically? Why aren't both ventricles included in the definition?
Does this mean that the left ventricle pumps more/less blood (through the aorta) than the right ventricle (through the pulmonary artery)?

Your reference gives the left and right ventricle stroke volumes, which are the same. For all practical purposes, it has to be the same in healthy individuals; if it were not, blood would back up somewhere and oxygenation would decrease (this does happen in pathological situations.)  Why the left? Because that's the definition of stroke volume. (Circular argument. I don't actually know why, because cardiac output can be measured easily now on both sides.) I imagine there's a better reason, but I'm not a cardiologist.

I’m voting to close this question because it is under researched, unclear (title and body don't match), and  contains multiple questions. These are all reasons for downvoting and closure on this site. Please take the [tour] and consult the [help] starting with [ask] for details and delete or [edit] accordingly.


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Rethinking SARS-CoV-1 and MERS-CoV deadliness after lessons learned from SARS-CoV-2
SARS-CoV-1 case fatality rates were about 9% and MERS-CoV case fatality rates were over 30%.
And now there is a newly discovered MERS-CoV relative, which some scientists have warned may be only one or two mutations away from making the jump to humans, and might be as deadly as MERS-CoV in humans.
But in the case of the SARS-CoV-2 pandemic, we've found that our initial CFR was higher than the true IFR. Can we expect that the CRFs for SARS-CoV-1 and MERS-CoV are similarly misleading? Are we able to estimate IFRs for these coronaviruses any better now that we have so much experience with SARS-CoV-2?

The CFR for SARS and MERS is calculated from the numbers of infected and died prople from the disease. Since both cause severe disease and as far as I know asymptomatic disease is not happening there (though I might be wrong) I don't think the numbers are way off.

If you have additional information relevant to your question please edit it into your post (comments are ephemeral and often overlooked). Adding sources to your question would make it stronger as would making the title more focused — as written it sounds like you are trying to start a discussion, which is expressly off-topic for this site. However, I encourage you to find sources in the medical literature or other reliable sources — the news media, even when they are trying to be honest, often oversimplify or make significant factual errors.

CFR is always higher (or at least as high) as IFR, since not every infection results in a symptomatic case. With SARS-CoV2, asymptomatic infections (or nearly asymptomatic one) are a big problem as they make contact tracing so difficult.

The reason CFR and IFR were off for this pandemic were that we had no idea of the full scale of the infections early on - pretty just those that turned up at hospitals and based on symptoms alone (i.e. no tests available). Once tests were developed and we could test anyone who was identified as a case or a contact, then the numbers rapidly fell and became more realistic.
Not so with SARS and MERS, where as far as we know most people get sick and were isolated so we can definitively say the CFR and IFRs. As SARS is no longer around, the CFR and IFR are known definitively.
Sero-prevalence (note this doesn't indicate infection necessarily, but exposure to the virus in some form) has been measured1 and found to be very low for MERS at around 15/10,000 in Saudi-Arabia. For SARS sero-prevalence was about 0.1%2 in asymptomatic people from those exposed to SARS-CoV-1 infected patients.
1: Muller. Lancet Infect Dis. 2015 May; 15(5):559-64.
2: Leung . Epidemiology & Infection, Volume 134, Issue 2, April 2006, pp. 211 - 221


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Simulate merging of two ssDNA together
I have two .pdb files with two ssDNAs, which should form some specific shape when closed together. Is it possible to model/simulate this merging somehow?
I would preferably like to have a program that would take both .pdb files as input and return .pdb file of the combined molecule.

If the structures are relatively small (< 4 nt), you can use Chem3D (from PerkinElmer ChemOffice software): Load both of the PDB files and run an MM2 minimization task.
For more complex structures GROMACS or similar molecular dynamics tools are used.